Hawkmoths of Australia

Maxwell Moulds; James Tuttle; David Lane

Hawkmoths are large charismatic insects with highly variable and colourful larvae. Some species are specialised in their habitat preferences, but others are widespread and often encountered in gardens. However, little is known about most species, and associating the adults with their larvae has previously been difficult or impossible. Hawkmoths of Australia allows identification of all of the Australian hawkmoths for the first time and treats species found on mainland Australia, Tasmania and all offshore islands within Australian limits. It presents previously undescribed life histories of nearly all species and provides a comprehensive account of hawkmoth biology, including new parasitoids and their hawkmoth hosts. Detailed drawings and photographs show the external and internal morphology of adults and immatures, and eggs, larval instars and pupa. Keys are provided for last instar larvae and pupae of the 71 species that the authors have reared. The book is concluded by a glossary, appendices to parasitoids and larval foodplants, an extensive reference list with bibliographical notes and a comprehensive index. The wealth of new information in this book makes it an essential reference for anyone interested in these moths. Hawkmoths of Australia is Volume 13 of the Monographs on Australian Lepidoptera Series.

Hawkmoths of Australia Identification, Biology and Distribution 190613 Hawkmoths of Australia 3pp.indd 1 29/08/19 11:10:58.97 Monographs on Australian Lepidoptera Editorial Board Editor-in-Chief Marianne Horak Australian National Insect Collection CSIRO NRCA GPO Box 1700 Canberra ACT 2601, Australia Editorial Panel M. F. Braby, Canberra, Australia E. D. Edwards, Canberra, Australia R. L. Kitching, Brisbane, Australia S. E. Miller, Washington DC, USA M. J. Scoble, London, UK M. S. Upton, Canberra, Australia Cizara ardeniae (Lewin, 1805), male Drawing by Sharyn Wragg 190613 Hawkmoths of Australia 3pp.indd 2 29/08/19 11:10:59.09 Monographs on Australian Lepidoptera Volume 13 Hawkmoths of Australia Identification, Biology and Distribution Maxwell S. Moulds, James P. Tuttle and David A. Lane 190613 Hawkmoths of Australia 3pp.indd 3 29/08/19 11:10:59.16 Dedication This book is dedicated to Ian Kitching who has so willingly shared his knowledge of hawkmoths with us and many others. © Maxwell S Moulds, James P Tuttle and David A Lane 2020 All rights reserved. Except under the conditions described in the Australian Copyright Act 1968 and subsequent amendments, no part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, duplicating or otherwise, without the prior permission of the copyright owner. Contact CSIRO Publishing for all permission requests. The authors assert their moral rights, including the right to be identified as an author. A catalogue record for this book is available from the National Library of Australia. ISBN: 9781486302819 (hbk) ISBN: 9781486302826 (epdf) ISBN: 9781486302833 (epub) How to cite: Moulds MS, Tuttle JP, Lane DA (2020) Hawkmoths of Australia: Identification, Biology and Distribution. Monographs on Australian Lepidoptera Volume 13. CSIRO Publishing, Melbourne. Published by: CSIRO Publishing Locked Bag 10 Clayton South VIC 3169 Australia Telephone: +61 3 9545 8400 Email: publishing.sales@csiro.au Website: www.publish.csiro.au Cover design by James Kelly Typeset by Desktop Concepts Pty Ltd, Melbourne Printed in China by Leo Paper Products Ltd. CSIRO Publishing publishes and distributes scientific, technical and health science books, magazines and journals from Australia to a worldwide audience and conducts these activities autonomously from the research activities of the Commonwealth Scientific and Industrial Research Organisation (CSIRO). The views expressed in this publication are those of the author(s) and do not necessarily represent those of, and should not be attributed to, the publisher or CSIRO. The copyright owner shall not be liable for technical or other errors or omissions contained herein. The reader/ user accepts all risks and responsibility for losses, damages, costs and other consequences resulting directly or indirectly from using this information. The paper this book is printed on is in accordance with the standards of the Forest Stewardship Council ® and other controlled material. The FSC® promotes environmentally responsible, socially beneficial and economically viable management of the world’s forests. Oct19_01 190613 Hawkmoths of Australia 3pp.indd 4 29/08/19 11:10:59.19 Contents Preface vii Acknowledgments viii Monographs on Australian Lepidoptera: previous volumes in this series Organisation and presentation Scope of the work Taxonomy Foodplant records The use of DNA sequence data for identification Plates Distributions Descriptions of adults and immature stages Taxonomic changes Abbreviations x 1 1 1 1 1 2 2 2 3 3 Historical review 4 Structure and function 6 Adult Egg Larva Pupa 6 14 14 15 Collection and preservation 19 Collecting adult hawkmoths Killing specimens Field storage Labelling specimens Preparing molecular specimens Dissecting genitalia The role of photography 19 19 20 20 20 20 21 Rearing hawkmoths Collecting immatures Eggs from wild-caught females Rearing larvae Artificial diets for larvae Housing pupae Rearing successive generations Biology Egg Larva Pupa Adult Hawkmoths as pests Hawkmoths as human food and medicine Natural enemies 190613 Hawkmoths of Australia 3pp.indd 5 22 22 22 22 23 23 23 24 24 24 25 26 27 27 27 29/08/19 11:10:59.22 vi HAWKMOTHS OF AUSTRALIA Classification and nomenclature 34 Higher classification Genus and species 34 34 The Australian fauna 36 Checklist of Australian species Key to last instar larvae Key to pupae Acherontia Acosmeryx Agrius Ambulyx Amplypterus Angonyx Cephonodes Cerberonoton Chelacnema gen. nov. Cizara Coenotes Coequosa Cypa Daphnis Eupanacra Eurypteryx Gnathothlibus Hippotion Hopliocnema Hyles Imber Leucomonia Macroglossum Megacorma Nephele Pseudoangonyx Psilogramma Synoecha Tetrachroa Theretra Zacria Addendum Daphnis Macroglossum Marumba 36 37 41 45 48 58 65 67 69 71 84 88 91 93 98 108 109 118 121 121 128 151 156 160 163 166 204 205 210 211 233 235 238 271 275 275 275 276 Plates 279 Glossary 371 Appendix 1: Sphingidae–Parasitoid associations 372 Appendix 2: Summary of known larval foodplants 376 References 382 Index 400 190613 Hawkmoths of Australia 3pp.indd 6 29/08/19 11:10:59.26 Preface The hawkmoths (Sphingidae) are among the largest and most showy of the world’s Lepidoptera. The adults are much sought after by collectors and the large larvae are often encountered by horticulturalists and gardeners. In spite of the hawkmoths’ somewhat high public profile, relatively little is known about the natural history of most species. Most of the historical knowledge about the Australian Sphingidae, as with all other regional sphingid faunas, has been focused primarily on the adults. This is not to suggest that some thoughtful and valuable research on the biology and ecology of the Australian hawkmoths has not been published; however, such publications tended to be narrow in their perspective. We entered into this project knowing that there were great gaps in the biological record of the Australian hawkmoths, and we understood the magnitude of the task that we had set for ourselves. Yet we believed that taxonomy would be a relatively simple matter for we rather naïvely assumed that the relationships of the Australian fauna were fairly well settled. However, it soon became obvious that a great deal of taxonomic work was also needed. How many species was such a vast continent hiding, either as outright clearly distinct taxa or unrecognised species concealed within species complexes? In an attempt to better understand the Australian sphingid fauna, both taxonomically and biologically, the authors pooled their knowledge about hawkmoths. MSM has 190613 Hawkmoths of Australia 3pp.indd 7 extensive field experience across much of Australia, including the many arid regions that are otherwise poorly sampled, and his reference collection served as an unparalleled starting point for our research. DAL has extensive field and rearing experience with the fauna of the rain forests of the Wet Tropics of north-eastern Queensland and the Northern Territory Top End. JPT is a newcomer to the Australian sphingid fauna but could draw on prior experience with the North American sphingids to transition into this research. All three authors have previously published on the Sphingidae and, in varying combinations, also on the Saturniidae and other Lepidoptera. We hope that this collaborative effort, with generous guidance from Ian Kitching from The Natural History Museum, London, will stabilise the taxonomic issues within the Australian sphingid fauna and answer many of the biological questions surrounding the various species. Yet as with all scientific endeavour, answering one question invariably leads to another. Understanding that limitation, it is our goal to establish a firm foundation upon which future researchers can build, hoping that this volume will inspire the next generation of sphingid workers. Maxwell S. Moulds James P. Tuttle David A. Lane 14 March, 2019 29/08/19 11:10:59.29 Acknowledgments We are particularly grateful to Marianne Horak (ANIC) and Ian Kitching (NHMUK) for critically reviewing the entire manuscript several times over many months. They provided many suggestions and corrections that have greatly improved this book. For comments on sections of the manuscript relating to their expertise, we thank Ted Edwards (ANIC), David Marshall (University of Connecticut), Roger Shivas (Department of Agriculture and Fisheries, Brisbane), Richard Weir (Department of Primary Industries and Fisheries, Darwin) and Andreas Zwick (ANIC). For carefully checking the references and other editorial matters we are very grateful to Murray Upton. For further technical advice, we thank the late Ian Common, Nikolaus Koeniger, Bert Orr and Rodolphe Rougerie. For generously providing important information from their personal experience, we thank Carol and Trevor Deane, Dominic Funnell, Bjørn Fjellstad, Denis Kitchin, Robert Lachlan, John Olive, Peter Mackey, Cliff Meyer, Garry Sankowsky, Thierry Salesne and Tom and David Sleep. Rodolphe Rougerie (Barcode of Life Data System project, BOLD) provided access to DNA barcodes which was invaluable in sorting out many of the taxonomic issues at the species level. Andreas Zwick (ANIC) and Daniel Rubinoff (University of Hawaii) similarly provided DNA barcodes. Akito Y. Kawahara (University of Florida), conducted multiple gene sequencing which was useful in assessing taxonomic issues at the generic level. To all, we express our sincere thanks. In addition, Tomás Melichar, Jean Haxaire and Ulf Eitschberger gave us permission to view their non-public DNA sequences within the BOLD system. Botanical assistance such as plant identifications and/or locating live plant material for rearing larvae was kindly provided by Jim Armstrong (Royal Botanic Gardens, Sydney), the late John Beasley (Kuranda), Sally Cowan (Australian Quarantine and Inspection Service), Neil Hoy (Rockhampton), the late Tony Irvine (Atherton CSIRO), Phil James (Eremophila Nursery of Western Australia), Stephen McKenna (Department of Environment and Resource Management), Chris Lane (Parks and Gardens, Shire of Broome), National Herbarium of New South Wales (Royal Botanic Gardens, Sydney), National Herbarium of Victoria (Royal Botanic Gardens, Melbourne), Peter North (Kuranda), Nick Smith (Rockhampton Council Nursery), Queensland Herbarium (Department of Environment and Resource Management), Garry and Nada Sankowsky (Tolga), Bruce Wannan (Department of Environment and Resource Management), Western Australia Herbarium (Department of Environment and Conservation), and Gary Wilson (Australian Tropical Herbarium). Foodplant records have been kindly provided by Graham Brown, the late Ian Common, Mike Daniell, Greg Daniels, Carol and Trevor Deane, Ted Edwards, Angus Emmott, Gary Fitt, Bjørn Fjellstad, Anne Garrett, the late Alan Graham, Tony Hiller, the late Sheila and Norman Hunter, the late Ailsa Johnson, Gordon Jones, Denis Kitchin, Robert Lachlan, the late Noel McFarland, Judy McMaugh, Geoff Monteith, Cliff Meyer, John Olive, Tony Rose, Garry Sankowsky, the late Courtenay Smithers, John Stockland, Bronwyn and Stephen Underwood, Peter Valentine, Maria and Allan WalfordHuggins and Geoff Williams. New foodplant records are 190613 Hawkmoths of Australia 3pp.indd 8 acknowledged individually in the foodplant list for each species. For identification of the varied natural enemies, we thank Art Borkent (Royal British Columbia Museum, Salmon Arm, British Columbia, Canada), Gavin Broad (Natural History Museum, London, England), Bryan Cantrell (Queensland Museum, Brisbane, Queensland, Australia), Peter Cranston (Australian National University, Canberra, ACT, Australia), José Fernández (University of Guelph, Guelph, Ontario, Canada), Gary Gibson (Canadian National Collection, Ottawa, Ontario, Canada), Henri Goulet (Canadian National Collection, Ottawa, Ontario, Canada), R. Bruce Halliday (ANIC, Canberra, ACT, Australia), John Heraty (University of California, Riverside, California, USA), Mike Hodda (ANIC, Canberra, ACT, Australia), John Huber (Canadian National Collection, Ottawa, Ontario, Canada), Lubo Masner (Canadian National Collection, Ottawa, Ontario, Canada), Donald Quicke (Chulalongkorn University, Bangkok, Thailand), Anthony Rice (Department of Agriculture and Water Resources, Cairns, Queensland, Australia), Stefan Schmidt (Zoologische Staatssammlung, Munich, Germany), and Ken Walker (Museum Victoria, Melbourne, Victoria, Australia). Living material of poorly understood or difficult to find species were provided by Sally Cowan, Carol and Trevor Deane, Bjørn Fjellstad, Dominic Funnell, John Olive, Wynne and Bruce Robinson, Garry Sankowsky, and Tom and David Sleep. For providing important leads in our field work that enabled us to complete life histories, we also thank the late John Beasley, Michael Braby, David Britton, Graham Brown, Glenn Cocking, Greg Daniels, Mike Gillam, Alan Graham, Tony Hiller, Jenny Holmes, Ernest Hoskin, David Knowles, Robert Lachlan, the late Noel McFarland, Stephen McKenna, Geoff Martin, Ian Moss, Geoff Monteith, Wendy Moore, Steve Morton, Craig Nieminski, Buck Richardson, Robert Richardson, Harley Rose, Anthony and Elizabeth Rice, Tony Rose, the late Courtenay Smithers, Alastair Stewart, Allen M. Sundholm, Allan and the late Maria Walford-Huggins, Bruce Wannan, Geoff Williams, Stephen and Bronwyn Underwood, and Peter Valentine. For field assistance, we thank Mark Lane in providing help in accessing remote areas throughout northern Australia, Ranger John Purdie for assistance with collecting in isolated areas of Katherine Gorge National Park and David Knowles for assistance with field work and equipment in Western Australia. In addition, MSM thanks Barbara and Timothy Moulds who accompanied him on numerous field trips across Australia over many years, to Kathy Hill and David Marshall for company on many others and to Margaret Humphrey for assistance in more recent field work throughout much of northern Queensland, Central Australia and New Guinea. We thank Buck Richardson for his considerable assistance in assembling the life history plates. The majority of those photographs were taken by the authors. For providing additional photographs used in the life history plates we are most grateful to: Hans Beste, Pls 6(n), 8(l), 16(m), 24(m), 29(n), 32(m), 61(k), 67(k), 68(m), 71(l); Sally Cowan, Pl. 35(m); Carol and Trevor Deane, Pls 24(g), 30(e, f), 31(m), 44(f), 67(i); Bjørn Fjellstad, Pls 12(g), 19(i), 20(k), 57(b, j); the late Alan Graham, Pl. 23(m); Melissa Harrison, Pl. 29(i); Jack and Sue Hasenpusch, 29/08/19 11:10:59.33 ACKNOWLEDGMENTS Pls 28(m), 31(h, i), 49(l); Alan Henderson, Pls 29(c), 50 (g), 67(f); Kathy Hill, Pl. 17(f, h); Jenny Holmes, Pl. 32(a); Margaret Humphrey, Pl. 64(h); Vivien Jones, Pl. 32(l); Sara Knight, Pl. 10(k); David Marshall, Pl 83(d); Cliff Meyer, Pl. 32(g); John Olive, Pls 14(d), 22(h), 49(g), 69(h); John Rainbird, Pl 83(c); David Rentz, Pls 4(c, h), 5(f, j), 7(a), 9(a, b, d, f, g), 14(a, b), 16(b, d, e, g), 17(a), 24(b, h), 26(b, i), 29(a), 30(i), 31(a), 32(b), 36(b, d), 42(d, i–l), 45(a, d, e, i, j), 46(d), 47(a-d, f-g, j), 50(d, h, j), 53(b, g, i), 57(f), 63(b, g), 67(c, e, j), 71(c); Buck Richardson, Pl. 58(j), Pl. 69(m); Garry Sankowsky, Pls 22(d), 23(g), 33(g), 41(l), 49(h); Josef Schofield and Danae Moore, Pl. 35(f, i); Tom Sleep, Pls 16(a), 52(l), 60(l), 62(k), 67(g), 68(i), 69(i); Rex Stock, 61(g); Allen M. Sundholm, Pl. 15(k); Stephen Underwood, Pl. 54(i); Gary Wilson, Pl. 63(f), Pl. 67(a); Paul Zborowski, Pl. 62(g). Photographs of male genitalia were taken by David Rentz, a huge task of several weeks for which we are most grateful. Microphotographs of parasitoids were taken by Clare McLellan and Simon Hinkley through the generous cooperation of Museum Victoria, Melbourne. Many other photographs for research or publication, including of adults, types, immatures and genitalia preparations, were provided by Hans Beste, Graham Brown, Ted Cadwallader, Glenn Cocking, the late Graeme Cocks, Sally Cowan, Greg Daniels, Carol and Trevor Deane, Fabian Douglas, Dominic Funnell, Ulf Eitschberger, Angus Emmott, Jennifer Ennion, Bjørn Fjellstad, Mike Gillam, the late Alan Graham, Mike Halsey, Jack and Sue Hasenpusch, Jean Haxaire, Alan Henderson, Kathy Hill, Jenny Holmes, Mark Hopkinson, Shane Hume, Margaret Humphrey, Paul Hutchinson, Jane Hyland, Vivien Jones, Paul Kay, Denis Kitchin, Ian Kitching, Sara Knight, Nikolaus Koeniger, Grant Kuseff, Robert Lachlan, Peter Marriott, Tomás Melichar, Cliff Meyer, Wendy Moore, Craig Nieminski, John Olive, Michael Powell, David Rentz, Buck Richardson, Wynne Robinson, Harley Rose, Tony Rose, Thierry Salesne, Garry Sankowsky, Tom and David Sleep, and Frank Standfast. In particular, we are especially grateful to Jean Haxaire, Robert Lachlan and Tomás Melichar for providing genitalia dissections from very rare species, critical to our study. Ivan Nozaic drew and inked the many line drawings used throughout this work. Ivan also provided translations from French. For both, we express our sincere thanks. This study would not have been possible without extensive specimen records. For providing specimens, distribution records or for helping with field work we are indebted to the following: Chris Ashhurst-Smith, Graham Brown, Steve Brown, Ted Cadwallader, Glenn Cocking, Sally Cowan, Greg Daniels, Fabian Douglas, Angus Emmott, Bjørn Fjellstad, Dominic Funnell, the late Alan Graham, Kees Green, Bart Hacobian, Mike Halsey, George Hangay, Jack and Sue Hasenpusch, Mark Heath, Peter Hendry, Cleave Herd, Marilyn Hewish, Kathy Hill, Tony and Kate Hiller, Jenny Holmes, Mark Hopkinson, the late Sheila Hunter, Paul Hutchinson, Ian Johnson, the late Steve Johnson, Denis Kitchin, Roger Kitching, John and Anne Koeyers, Mark Lane, Rob Lachlan, the late Noel McFarland, Peter Mackey, Dave Marshall, Peter Marriott, Cliff Meyer, Geoff Monteith, Wendy Moore, Chris Müller, Sue and Michael Murphy, Craig Nieminski, Ivan Nozaic, Erica Odell, John Olive, Michael Powell, Clive Pratt, David Rentz, Anthony Rice, Buck Richardson, Wynne Robinson, Tony Rose, Don Sands, Alastair Stewart, Allen Sundholm, Ken Thomsen, Bronwyn and Stephen Underwood, Peter Valentine, Allan Walford-Huggins, James Walker, Geoff Williams, Steve Williams and Theo Wright; together they have contributed numerous important specimen records over many years. Special thanks to Dion Maple and Caitlyn Pink 190613 Hawkmoths of Australia 3pp.indd 9 ix (Parks Australia, Christmas Island) for not only collecting on our behalf for two years, but facilitating the internal permit process and importation issues. We are also grateful to the curators and staff of the following collections for providing information on specimens in their care or for providing access to their collections: ANIC (Ted Edwards, You Ning Su, Marianne Horak and the late Ebbe Nielsen); AM (David Britton, Derek Smith and John Tann); NHMUK (Geoff Martin and Ian Kitching); CMNH (John Rawlins and Jane Hyland); MCA (Adam Yates); MM (Margaret Humphrey); MV (Peter Lillywhite, Simon Hinkley, Catriona McPhee, Peter Marriott and Ken Walker); NTM (Michael Braby and Gavin Dally); QM (Geoff Monteith, Christine Lambkin and Chris Burwell); QVM (Lisa Gershwin); RNH (Willem Hogenes); SAM (Jan Forrest and Peter Hudson); TMAG (Catherine Byrne); WADA, Perth (Andras Szito); WADA, Kununurra (Geoff Strickland); WAM (Terry Houston and Brian Hanich). In particular, Ted Edwards, Brian Hanich, Peter Hudson, Ian Kitching, Peter Lillywhite, Simon Hinkley and You Ning Su are especially thanked for providing photographs and answering our many questions relating to material in their collections. MSM thanks Barbara Moulds for numerous hours of collection curation over many years. For testing the keys to species for larvae, pupae and Psilogramma adults, we thank Glenn Cocking, Sally Cowan, Dominic Funnell, Marianne Horak, Margaret Humphrey, Paul Hutchinson, Jean Weiner and Gary and Robyn Wilson. For copies of literature or access to their libraries, we are indebted to the librarians and staff of the Australian Museum, Sydney; Museum Victoria, Melbourne; Natural History Museum, London; and CSIRO Black Mountain Laboratories, Canberra. Further, we wish to thank Ron Brechlin, Hans Duffels, Ted Edwards, Ulf Eitschberger, Jean Haxaire, Roger Kendrick, Ian Kitching, Tomás Melichar, James O’Hara and Werner Schmidt (ZAG Wirbellose e. V. Publications) for copies of difficult-to-access literature. For access to Hammond Island, Torres Strait, we are grateful for the generous support of Councillor Mario Sabatino of Torres Strait Island Regional Council and Rita Dorante, Brian Arndt and Sharon Sabatino for providing accommodation and much assistance during our several visits. Similarly, for permission to visit and collect on Dauan Island, Torres Strait, we thank Councillor Torenzo Elisala and the Elders of Dauan Island, and we acknowledge the support of Liz and Wayne Phillip in providing invaluable assistance during our visits. Without their generosity and help our visits to Torres Strait would not have been possible. For collecting permits, we thank the NSW National Parks and Wildlife Service; Forestry Commission of NSW; National Parks, Northern Territory; Department of Forestry, Queensland, National Parks and Wildlife Service, Queensland, and Department of Conservation and Land Management, Western Australia. In particular, we are grateful to the Queensland Entomological Society for administering the provision of the Queensland National Parks Collecting Permits and to the National Parks field staff for their support. Special thanks are extended to Richie Carrigan, Senior Ranger at Wooroonooran National Park, for his advice and assistance. For financial assistance, we thank the Australian Museum and the Linnean Society of New South Wales for a Joyce Vickery Research Award. Without this assistance, our project would have been compromised. Much of the manuscript was typed by Sally Cowan and Barbara Moulds; we thank both for their many hours of work. 29/08/19 11:10:59.38 Monographs on Australian Lepidoptera: previous volumes in this series Volume 1, 1989 Primitive Ghost Moths Morphology and Taxonomy of the Australian Genus Fraus Walker (Lepidoptera: Hepialidae s. lat.) E.S. Nielsen and N.P. Kristensen Volume 8, 2000 Oecophorine Genera of Australia III The Barea Group and Unplaced Genera (Lepidoptera: Oecophoridae) I.F.B. Common Volume 2, 1993 Tineid Genera of Australia (Lepidoptera) G.S. Robinson and E.S. Nielsen Volume 9, 2004 Zygaenid Moths of Australia A Revision of the Australian Zygaenidae (Procridinae: Artonini) Gerhard M. Tarmann Volume 3, 1994 Oecophorine Genera of Australia I The Wingia Group (Lepidoptera: Oecophoridae) I.F.B. Common Volume 4, 1996 Checklist of the Lepidoptera of Australia E.S. Nielsen, E.D. Edwards and T.V. Rangsi (Editors) Volume 5, 1997 Oecophorine Genera of Australia II The Chezala, Philobota and Eulechria Groups (Lepidoptera: Oecophoridae) I.F.B. Common Volume 6, 1999 Biology of Australian Butterflies R.L. Kitching, E. Scheermeyer, R.E. Jones and N.E. Pierce (Editors) Volume 10, 2006 Olethreutine Moths of Australia (Lepidoptera: Tortricidae) Marianne Horak with contributions by Furumi Komai Volume 11, 2011 Elachistine Moths of Australia (Lepidoptera: Gelechioidea: Elachistidae) Lauri Kaila with contributions by Kazuhiro Sugisima Volume 12, 2018 Splendid Ghost Moths and Their Allies A Revision of Australian Abantiades, Oncopera, Aenetus, Archaeoaenetus and Zelotypia (Hepialidae) Thomas J. Simonsen Volume 7, 1999 Heliothine Moths of Australia A Guide to Pest Bollworms and Related Noctuid Groups M. Matthew 190613 Hawkmoths of Australia 3pp.indd 10 29/08/19 11:10:59.42 Organisation and presentation While this book is written with the hawkmoth enthusiast in mind, we hope that it will be useful across a broad level of interests from the professional entomologist to the general naturalist. Scope of the work We treat all species found through mainland Australia, Tasmania, and all offshore islands within Australian limits including the islands of the Torres Strait, the islands of the Great Barrier Reef, Lord Howe Island, Norfolk Island, Christmas Island (Indian Ocean) and the Cocos-Keeling Islands (Fig. 1). Taxonomy The detailed treatment of the Australian fauna is arranged alphabetically by genera then alphabetically by species within genera. Our Checklist gives the taxa listed according to the current classification but the higher classification of hawkmoths requires further study with many unresolved issues. Names of species and subspecies not found in Australia follow current literature. We follow Kitching and Cadiou (2000) in using original spelling of species names even though some do not agree in gender with their generic combination as required by the International Code of Zoological Nomenclature, third edition. This is a convention now widely accepted within Lepidoptera circles that engenders stability for electronic searches. When listing synonymies, we include not only names that are junior synonyms but all different generic combinations of these names, together with the reference to the first use of these combinations. Foodplant records This work contains a large number of new foodplant records, most originating with the authors. The majority of previously published records have been confirmed and erroneous records have been noted and excluded from Appendix 2. New foodplant records not originating from us are explicitly acknowledged and referenced. Plant names follow those used by the Census of the Queensland Flora 2017 and the Australian Plant Census (APC). Exotic plant names not listed in the above sources follow those of the World Checklist of Selected Plant Families (WCSP). Plant names preceded by an asterisk (*) indicate nonnative species. The use of DNA sequence data for identification An increasingly popular method for species identification is the matching of DNA sequence data against a database of reference sequences, which is commonly referred as ‘DNA Fig. 1. Geographic region covered by this book. 190613 Hawkmoths of Australia 3pp.indd 1 29/08/19 11:11:00.30 2 HAWKMOTHS OF AUSTRALIA barcoding’. For Lepidoptera, a 658bp fragment of the mitochondrial gene cytochrome C oxidase subunit 1 (COI) has been established as a standard marker that is relatively easy to amplify and sequence, even from dried and old specimens. The Barcode of Life Data Systems project (BOLD) is the largest publically accessible reference database that hosts userprovided identifications, images and sequence data for many different organisms worldwide. Although the library is far from complete, it provides a framework for addressing many taxonomic questions. As of this date, BOLD holds 23 000+ DNA sequences of Sphingidae, 1200+ of which are of Australian origin (Rougerie et al. 2014). In addition to the data submitted by others, we added many Australian samples to fill gaps in the taxonomic and geographic coverage, the majority of which were derived from adults reared from larvae. Many of those barcoded adults are illustrated herein and linked to BOLD by their unique sample identification number (BC-LTM-***). We used DNA barcode sequences in BOLD in several ways, primarily for comparing taxa, assessing allopatric populations within species complexes and genetic diversity within single populations that exibited distinct morphological variation. As a simple, subjective ‘rule of thumb’, we consider a greater than 2.0% genetic distance (difference in DNA sequences) between an individual and its nearest match to be suggestive of those individuals belonging to distinct species. A more sophisticated approach is the use of clustering algorithms as implemented in the Barcode Index Number (BIN) system (Ratnasingham and Hebert 2013) of BOLD. In our case of Australian Sphingidae, DNA barcode results usually matched long-held species concepts, but sometimes also revealed the potential presence of currently unrecognised species, including undescribed species. The most striking example of how DNA barcodes helped us identify adults in a difficult species complex and associate their larvae was in the genus Psilogramma. Prior to the checklist of Moulds (1996), who listed two species from Australia, only one Psilogramma species (P. menephron) was recognised from Australia and all larval identifications were attributed to that single species. Three years later, Moulds and Lane (1999) described a third species, P. argos, designating a reared specimen as holotype. In doing so, they created the first verifiable larval-adult Psilogramma association from Australia. Two years later, Brechlin (2001) and Eitschberger (2001a, 2001b) described several additional Psilogramma species and revised the status of others bringing the total number of names associated with the Australian Psilogramma fauna to ten. Subsequent barcoding (Rougerie et al. 2014) and taxonomic analysis has reduced that number, as recognised herein, to eight. However, with the exceptions of P. argos and the resurrected P. casuarinae, whose distribution extends much further south than any other Psilogramma species, thereby eliminating confusion in Victoria and much of New South Wales, the Australian Psilogramma are not only interspecifically similar in appearance but also intraspecifically variable in their markings. We have now reared and fully documented the life histories of six of the eight Psilogramma species and had the COI barcode region of multiple reared adults sequenced. By using DNA barcodes we were able to obtain positive adult identifications enabling us to work out how to identify adults from their markings and also to correctly identify and associate larvae and their foodplants. However, relying upon short sections of a single gene has its limitations and exclusive reliance upon DNA barcodes can be misleading. As an example at one extreme, our submissions of Hippotion boerhaviae and H. rosetta to BOLD had a minimum genetic distance of only 0.46% and, on the face of it, tended to support a previously held belief that H. rosetta did 190613 Hawkmoths of Australia 3pp.indd 2 not occur in Australia. However, our H. rosetta and H. boerhaviae submissions were not based upon wild collected adults that could be potentially misidentified but rather upon adults confirmed through rearing; the larvae of H. rosetta and H. boerhaviae are markedly different in all five larval instars. By sequencing adults bred from known larvae, we were able to recognise the small 0.46% genetic distance as misleading, to confirm that H. boerhaviae and H. rosetta are indeed distinct species, and that both occur in Australia. At the other extreme, Theretra nessus has a vast geographical range that extends from Australia and some Melanesian islands across much of Asia west to the Indian subcontinent. Comparing a large number of Asian T. nessus DNA barcodes in BOLD, the smallest genetic distance to Australian specimens of T. nessus is 3.38% (Rougerie et al. 2014). In spite of this significant genetic variation, we were unable to find a single morphological or ecological character in the adults or larvae that would allow us to distinguish between populations. As a result, without additional evidence beyond the DNA barcode, we take a cautious approach and do not treat T. nessus as two species. Plates In our images we present the larva upright, apparently sitting on the upper leaf surface, a position seldom observed in nature. This is a deliberate approach, based on our assumption that one is more familiar with an upright caterpillar’s morphology and hence can more easily compare diagnostic characters in this orientation. Adult plates attempt to show the range of variation in variable species but the reader should be aware that intermediate or additional variants are likely. This is particularly relevant in genera such as Psilogramma where variation in wing pattern is sometimes extensive with almost no two individuals identical and the range of variation overlapping species boundaries. The plates of the male genitalia illustrate a typical example of each species as far as that is possible. The valvae have been spread to expose their inner surfaces and to show diagnostic features to their best advantage, and the tegumen carrying the uncus and gnathos has been cut free on one side to allow it to turn sideways for best viewing. It is important to note that there is often some variation between individuals even in diagnostic features and sometimes there may be minor distortion in structures due to positioning. It is strongly recommended that the genital descriptions be consulted for correct interpretation. Distributions Distribution records have been based only on specimens for which we have personally confirmed species identification. Significant distribution records from private collections (other than the authors’) are each acknowledged by inclusion of the collector’s name with the locality in the text. Descriptions of adults and immature stages Forewing measurements are provided as a guide to the size of the species and include the smallest and largest wild-caught individuals known to us. We do not attempt to differentiate the sizes of the sexes unless there is a significant difference between them. Proboscis measurements are the shortest and longest encountered, usually based on five individuals by attempting to include the smallest and largest specimens. The reader should be aware that larval descriptions may not necessarily encompass the full range of colour variability in a species because there may be variants we have not seen. Similarly, measurements included with the descriptions of eggs and larval instars may not always comprise the full size range. 29/08/19 11:11:00.37 ORGANISATION AND PRESENTATION When available, the number of days as egg and larval instars are given. These are not absolute figures and may depend on external factors such as temperature, type and condition of the foodplant, and humidity. Taxonomic changes The following taxonomic changes are proposed in this book: Acosmeryx cinnamomea (Herrich-Schäffer, 1869) stat. rev. is removed from synonymy with Acosmeryx anceus. Cephonodes australis Kitching and Cadiou, 2000 stat. nov. is given specific rank, previously considered a subspecies of Cephonodes hylas australis. Cephonodes cunninghami (Cramer, 1777) stat. nov. is removed from synonymy with Cephonodes picus (Cramer, 1777). Cerberonoton severina (Miskin, 1891) stat. rev. is returned to specific rank previously considered a subspecies of Cerberonoton rubescens. Chelacnema Moulds, Tuttle and Lane gen. nov. is described as new. Chelacnema ochra (Tuttle, Moulds and Lane, 2012) comb. nov. is transferred to Chelacnema from Hopliocnema Rothschild and Jordan, 1903. Hippotion johanna (Kirby, 1877) stat. rev. is reinstated to specific rank from being considered a form of Hippotion brennus (Stoll, 1782). Macroglossum corythus approximans (T.P. Lucas, 1891) stat. nov. is removed from synonymy with Macroglossum corythus pylene C. Felder, 1861. Macroglossum errans Walker, 1856 stat. nov. is given specific rank, previously considered a subspecies of Macroglossum hirundo Boisduval, 1832. Macroglossum melas pullius Jordan, 1930 syn. nov. is synonymised with Macroglossum melas melas Rothschild and Jordan, 1903. Macroglossum papuanum Rothschild and Jordan, 1903 stat. nov. is given specific rank, previously considered a subspecies of Macroglossum troglodytus Boisduval, [1875]. Macroglossum queenslandi Clark, 1927 stat. nov. is given specific rank, previously considered a subspecies of Macroglossum divergens Walker, 1856. Macroglossum stenoxanthum Turner, 1925 syn. nov. is synonymised with Macroglossum corythus approximans (T.P. Lucas, 1891). Theretra clotho manuselensis Joicey and Talbot, 1921 syn. nov. is synonymised with Theretra indistincta indistincta (Butler, 1877). Theretra clotho papuensis Joicey and Talbot, 1921 syn. nov. is synonymised with Theretra indistincta indistincta (Butler, 1877). Theretra latreillii prattorum Clark, 1924 syn. rev. is synonymised with Theretra lucasii (Walker, 1856). Theretra lucasii (Walker, 1856) stat. rev. is given specific rank, previously considered a subspecies of Theretra latreillii (W.S. Macleay, 1826). Theretra oldenlandiae firmata (Walker, 1856) syn. nov. is synonymised with Theretra oldenlandiae oldenlandiae (Fabricius, 1775) Theretra oldenlandiae lewini (Thon, [1828]) syn. nov. is synonymised with Theretra oldenlandiae oldenlandiae (Fabricius, 1775). Theretra oldenlandiae samoana syn. nov. is synonymised with Theretra oldenlandiae oldenlandiae (Fabricius, 1775). 190613 Hawkmoths of Australia 3pp.indd 3 3 Abbreviations The abbreviations used in the text are as follows: AM – Australian Museum, Sydney ANIC – Australian National Insect Collection, Canberra AR – Anthony Rice BMF – Bjørn M. Fjellstad NHMUK – Natural History Museum, London, UK BPBM - Bernice P. Bishop Museum, Hawaii CMNH – Carnegie Museum of Natural History, Pittsburgh, Pennsylvania, USA CN – Craig Nieminski CTD – Carol and Trevor Deane CV – cultivar DAL – David A. Lane DR – David Rentz EME – Entomologisches Museum Eitschberger, Marktleuthen, Germany GS – Garry Sankowsky GWW – Gary W. Wilson HAX – Jean Haxaire JH – Jack Hasenpusch JO – John Olive JPT – James P. Tuttle loc. – locality MCA – Museum of Central Australia, Alice Springs MM – Macleay Museum, University of Sydney MSM – Maxwell S. Moulds MV – Museum Victoria, Melbourne NS – Nada Sankowsky NTM – Northern Territory Museum, Darwin QM – Queensland Museum, Brisbane QVM – Queen Victoria Museum, Launceston RBL – R.B. Lachlan SAM – South Australian Museum, Adelaide SFV – Stephanus (Fanie) Venter SMCR – Sphingidae Museum, Czech Republic TDS – Tom and David Sleep TM – Tomas Melichar TMAG –Tasmanian Museum and Art Gallery, Hobart WADA – Western Australian Department of Agriculture and Food WAM – Western Australian Museum, Perth ZSM – Zoologische Staatssammlung, Munich, Germany 29/08/19 11:11:00.46 Historical review The earliest observations of Australia’s sphingid fauna were made through the lens of European experience. Pittaway (1993) provides an excellent account of the evolving published documentation of the hawkmoths of England and Europe that began in 1589 with the naming and the illustrations of the adult and larva of porcellus (a name subsequently adopted by Linnaeus for Sphinx porcellus = Deilephila porcellus). Among the works noted by Pittaway (1993), one deserves special attention as it would later define the name of the family. Réaumur (1736: xlvi) illustrated the larva, pupa, and adult of the Privet Hawk (Sphinx ligustri Linnaeus) and commented that the resting posture of the larva suggested the Sphinx of Egyptian antiquity. When Linnaeus (1758) first universally applied binomial nomenclature to zoology in the Systema Naturae, he selected Réaumur’s ‘Sphinx’ as the name for the genus that included 16 European hawkmoths and a number of additional taxa that were subsequently re-assigned to other families. Accordingly, Latreille [1802] named the hawkmoth family Sphingides. Bringing the Sphingidae into the modern era, Butler (1876b) removed all taxa that did not fit our current definition of the hawkmoths. Specifically regarding the Australian sphingid fauna, the two cosmopolitan species Agrius convolvuli and Hippotion celerio were among the 16 European sphingids named by Linnaeus (1758) in the 10th edition of his Systema Naturae. Quite ironically, these two well-known members of the Australian sphingid fauna were named over 10 years before Captain Cook’s arrival in Australia in 1770. Not surprisingly, the earliest known descriptions and illustrations of Australian sphingids were published in Europe (Donovan 1805; Lewin 1805; Perry 1811; W.S. Macleay 1826; Boisduval 1832; Angas 1847). The 1805 publications of both Donovan and Lewin were books containing beautiful hand-coloured plates, and they were the first published records of hawkmoths specifically from Australia. By the middle of the 19th century as the British Museum and the Museum of the Royal Dublin Society increased their holdings, most of the descriptions of new Australian Sphingidae resulted from material held in the British Isles and appeared mostly in the various scientific publications in London (Walker 1856, [1865]; Butler [1876a], 1877a; Kirby 1877). The one notable exception was the description of Darapsa moorei published by W.J. Macleay in 1866 in volume 1 of the Transactions of the Entomological Society of New South Wales, the first Australian entomological journal. Sir William Macleay, resident in Sydney, was an ardent collector of insects and instrumental in establishing the Entomological Society. By the 1890s several Australian entomologists began to self-publish and/or publish in Australian journals, and the documented sphingid fauna increased significantly (e.g. Olliff 1890; Lucas 1891a, 1892; Miskin 1891; Scott 1890–98; Lidgett 1893; Lower 1897a). Principal among these was Miskin’s (1891) synoptic treatment of the Australian Sphingidae then comprising 45 known species. Lucas and Miskin both resided in Brisbane and were arch rivals in the world of Australian Lepidoptera (Moulds 1999). Their race to publish descriptions of new species in 1891 and their isolation from the large European reference collections led to the establishment of ten 190613 Hawkmoths of Australia 3pp.indd 4 junior synonyms, although nine of their species have survived the test of time. Kirby (1892) catalogued the Sphingidae of the world and corrected several errors in Miskin’s (1891) synoptic treatment. In the same year, Swinhoe (1892) published his catalogue of Eastern and Australian moths in the collection of the Oxford University Museum, describing several new species in the process, including two from Australia, one of which is now a junior synonym. However, it was Rothschild and Jordan’s (1903) classic monograph running to almost 1000 pages [and the updated précis of this work in Genera Insectorum (Rothschild and Jordan 1907)] that created a classification that stabilised sphingid systematics and is largely accepted to this day. Their monograph described four new Australian species, one new Australian subspecies, and addressed several higher classification issues by creating six new Australian genera. Soon afterwards, Wagner (1913–19) published his checklist of the sphingids of the world with exhaustive literature citations. Subsequently, the list of Australian Sphingidae remained nearly unchanged for almost 80 years with only two subspecies added (Clark 1922, 1927). In the last two decades of the 20th century two new species were added (Moulds 1983; Moulds and Lane 1999), and Lachlan (1988) recorded two species previously unknown from Australia. Comprehensive treatments that addressed the Australian sphingid fauna during the latter 20th century were D’Abrera’s [1987] Sphingidae Mundi, illustrating almost every known species worldwide in colour together with associated text, Common’s (1990) Moths of Australia, providing an overview of the Australian hawkmoth fauna, and Moulds’ (1985) review of the Australian species of Macroglossum. Bridges (1993) and Moulds (1996) presented the first annotated synoptic checklists in almost 100 years. In addition, there has been a number of notable works that include sphingids whose distributions extend into Australia. Among the more significant are the monographic treatments of Tutt (1904), Mell (1922), Bell and Scott (1937), Dupont and Roepke (1941), Pinhey (1962), Holloway (1976), Pittaway (1993) and Danner et al. (1998). Throughout the 19th and 20th centuries little attention had been paid to the biology of Australian hawkmoths and, for the most part, only relatively brief accounts were recorded. Lewin (1805) was the first to document the early stages of Australian species when he depicted the larvae, pupae and foodplants of Cizara ardeniae (Lewin) and Theretra oldenlandiae (Fabricius), together with brief notes. It was over 80 years later that Tepper (1888) provided notes on the larvae and natural history observations on Hippotion celerio and H. scrofa (as Chaerocampa pallicosta (Walker)), which he repeated, almost verbatim, just two years later (Tepper 1890). Around this time, Scott (1864, 1890–98) published his Australian Lepidoptera and their transformations, remarkable for its exquisite plates of moths and butterflies with their life histories painted by his daughters, Harriet and Helena. However, only one of these plates depicted a hawkmoth even though Harriett and Helena had prepared several others (including three of hawkmoths) that remained unpublished until Ord (1988) reproduced them for the first time in a volume 29/08/19 11:11:00.53 HISTORICAL REVIEW on the Scott sisters. For the next 100 years, only short accounts concerning biology and distribution appeared, almost all as part of broader treatments of other groups; the few that related solely to Australian hawkmoths were only brief notes. These are far too numerous to list here and none stand out sufficiently to warrant special discussion. The many foodplant records scattered through the literature were eventually brought together, along with many new foodplant records, by Moulds (1981, 1984, 1998), but perhaps the most significant aspects of the first two of these papers were the figures of the larvae of 13 species and a larval key to 21 species respectively. Groth (1995) described the life history for Coequosa australasiae (Donovan), the most detailed account for any Australian hawkmoth to that date. More recently, new standards have been set in documenting the biology of Australian sphingids and four new life histories have been recorded; Lane (2006, 2009) described the life histories of Leucomonia bethia and Macroglossum prometheus lineata Lucas respectively, Lane and Moulds (2010) that of Imber tropicus Moulds (as Langia tropicus) in unprecedented detail, and Hasenpusch, Lane and Moulds (2012) similarly documented the life history of Macroglossum papuanum (as M. insipida). Kitching and Cadiou’s (2000) annotated revisionary checklist of the world’s Sphingidae offered a clear demarcation between the past and the present, not only by the heralding of a new millennium but by considering and integrating almost 100 years of disparate information into their taxonomic decision making. Papers describing new Australian sphingids since Kitching and Cadiou include Brechlin (2001) and Eitschberger (2001a, 2001b), two of which were published almost simultaneously (with the inevitable synonymy), and describing between them a remarkable 43 new species raising the number of named Australian Psilogramma from two to eight (although some of these names were later found to be synonyms). Further studies of Psilogramma (Eitschberger 2004a, 2010a, 2010b; Brechlin and Kitching 2010b; Lane, Moulds and Tuttle 2011), brought the number of Australian Psilogramma species to nine. In addition, Haxaire and Melichar (2003), Lachlan (2004b), Eitschberger (2010c), 190613 Hawkmoths of Australia 3pp.indd 5 5 Moulds, Tuttle and Lane (2010), Tuttle, Moulds and Lane (2012), Zolotuhin and Ryabov (2012), and Moulds and Melichar [2014] described five further new species (Chelacnema ochra, Coenotes arida, Gnathothlibus australiensis, Hopliocnema lacunosa, Zacria vojtechi) and four new genera (Cerberonoton, Imber, Pseudoangonyx, Zacria) from Australia. Although no further taxa have been described from Australia since 2014, Kitching et al. (2018b) made many changes to the status of taxa, mainly through synonymy. This had a relatively minor impact on the number of recognised species in Australia (it reduced the number of recognised species by just two) but it did affect the names of several taxa. Herein, we make further changes, including the addition of eight species to the Australian fauna (four only known from Christmas Island), and the description of the new genus Chelacnema. Recent progress in sphingid phylogenetics has been in large part due to advances in DNA analyses. Hundsdoerfer, Tshibangu et al. (2005), Kawahara et al. (2009) and Kawahara and Barber (2015) published molecular phylogenies incorporating some Australian species, complementing the earlier ground-breaking morphological phylogenies by Kitching (2002, 2003) that also incorporated Australian species. The broader finding of the studies by Kawahara et al. (2009) and Kawahara and Barber (2015), as they relate to systematics, are discussed below under ‘Classification and nomenclature–Higher classification’, and a revised higher classifcation based primarily on these studies was published by Kitching et al. (2018a). The study by Kawahara and Barber (2015), orientated towards the evolution of sphingid hearing, ultrasound production and bat sonar jamming, also provided palaeontological dating for the nodes on their tree. Kamaluddin et al. (1999, 2014) attempted intuitive phylogenies for the sphingids of Pakistan and Azad Kashmir using morphological characters. Kawahara et al. (2009) also provide a detailed historical account of the development of sphingid phylogenetics. A molecular study of the Lepidoptera as a whole by Breinholt et al. (2018) confirmed the sister relationship of the Sphingidae with the Saturniidae. 29/08/19 11:11:00.59 Structure and function Notable accounts of adult sphingid morphology can be found in Rothschild and Jordan (1903) and Kitching and Cadiou (2000). Kristensen (2003a, 2003b) gives a comprehensive account of adult Lepidoptera morphology but for the most part does not specifically mention sphingids. Common (1990) provides information on both general Lepidoptera morphology and specifics for sphingids. Stehr (1987) and Hasenfuss and Kristensen (2003) give detailed overviews of the morphology of the immature stages. Adult Head (Figs 2–5) The head is more or less rounded and always densely scaled, with the dorsal region referred to as vertex and the anterior part as frons. The head is dominated by the large compound eyes, a pair of long robust antennae and very often a long proboscis that is tightly coiled at rest. Eyes (Figs 2, 4, 5). Typical for insects, the large compound eyes are made up of numerous hexagonal facets or ommatidia. Hawkmoth eyes are known to have as many as 30 000 ommatidia. In some species the eyes are lashed dorsally and sometimes also ventrally by long setae. Ocelli and chaetosemata are lacking in hawkmoths. Yagi and Koyama (1963) describe unusual branched processes on the ring-shaped ocular apodeme supporting the compound eye that may be unique to hawkmoths. Detailed accounts of eye structure in hawkmoths can be found in Eguchi (1982) and Warran et al. (1999). Antennae (Fig. 5). The antennae are primarily sensory organs for smell but also play important roles in sensing orientation, gravity, temperature, humidity and air flow. They are usually a little shorter than half the forewing length and show at least subtle differences between the sexes. The basal segment is known as the scape, the second segment as the pedicel and a distal multi-segmented section is called the flagellum (Fig. 5). The pedicel carries a cluster of sensilla known as Johnston’s organ (Van den Berg 1971; Sane et al. 2007) that aids in perceiving movement and provides stability during flight. The flagellum of male hawkmoths has a characteristic structure with the ventral, unscaled surface of each segment (flagellomere) laterally concave and with long sensory cilia along both the anterior and posterior margins. Some females have a similar structure but their antenna is usually pubescent ventrally. In both sexes the flagellum is usually filiform but in the males of some genera, and in the females of one genus, none of which are found in Australia, the flagellum is bipectinate (Kitching and Cadiou 2000). In all but diurnal species the distal part of the flagellum tapers noticeably and the apical portion is recurved in a hook-like manner. In the diurnal Cephonodes the antennae are somewhat thickened distally. Some further accounts of antennal function and structure are given by Hallberg et al. (2003), Hinterwirth and Daniel (2010) and Nirazawa et al. (2017). Mouthparts and feeding (Figs 2–5). The proboscis (Figs 2, 5) is used for feeding, mostly for extracting nectar from flowers although a few tropical species take exudates from the eyes of animals and Acherontia species steal honey from bees of the genus Apis. In most hawkmoths it is well developed, often reaching lengths exceeding that of the body. In some 190613 Hawkmoths of Australia 3pp.indd 6 species such as Agrius convolvuli it can exceed 100 mm and in the Neotropical Amphimoea walkeri it reaches an extreme length approaching 300 mm (figured in D’Abrera [1987]). However, in about one-fifth of hawkmoths worldwide it is very short (Miller 1997). In some Australian Smerinthini including Coequosa and Cypa it may be functionally limited and in some Sphingulini, including Hopliocnema, it is so reduced as to be effectively absent. Needless to say, species without a proboscis cannot feed and their adult life lasts no more than 10 days to a fortnight, whereas species that feed can live six weeks or more. The proboscis is tightly coiled at rest but fully extended when feeding, and always with a distinct bend before midlength, the purpose of which is not fully understood. How the proboscis is extended and recoiled is complex, but essentially extension is achieved by an increase in haemolymph pressure in association with muscle movement while recoil is mainly a natural return following reduction in haemolymph pressure and muscle relaxation (Wannenmacher and Wasserthal 2003). The proboscis is formed from the paired elongated galeae of the maxillae, the inner surfaces of which are concave and when interlocked form the tubular food canal. Especially towards its tip it is covered with sensory organs for smell and taste for finding and evaluating food. A detailed account of proboscis structure and function is provided by Kristensen (2003a) and of proboscis musculature by Wannenmacher and Wasserthal (2003). Flanking the base of the proboscis are the paired labial palps (often just referred to as palps) that also are structurally part of the mouthparts. In hawkmoths these are 3-segmented, with the apical segment very small and often concealed within the distal scales of the penultimate segment. Rothschild and Jordan (1903) found the distribution of scales on the inner surface of the labial palps useful in diagnosing some tribes and genera, although they did not realise the differences were related to sound reception (see Hearing below). The maxillary palps are very small and reduced to a single segment barely discernible. Also at the base of the proboscis, above the palps, are the pilifers, a pair of very small, lobe-like, sensory structures. Unique to Acherontia, the mouthparts are also used in sound production. Both sexes can produce squeaking sounds that differ between species (Kitching 2003) and are produced by air movement through the proboscis. These sounds have twin origins, initially by air drawn in through the proboscis via a dilated pharynx which produces a rapid train of pulses, followed by expelled air that produces a brief sustained sound (Busnel and Dumortier 1959; Brehm et al. 2015). In A. atropos the process lasts only about 200 milliseconds and is repeated some 40–50 times to create a complete audible squeak. Hearing and palps (Figs 2–5). There are no hearing organs on the abdomen or thorax in hawkmoths unlike in many other larger moths. However, some species have modifications to segment 2 of the labial palps (Fig. 3) and the adjacent pilifer for detecting ultrasonic sound. The ability to perceive ultrasonic sound helps in avoiding predatory bats (Kawahara and Barber 2015; Hofstede and Ratcliffe 2016). Sound is received by the tympanum-like palps and the vibrations transferred to bristles on the adjacent pilifers and 29/08/19 11:11:00.67 STRUCTURE AND FUNCTION thence to the brain. Such a system is found in the subtribe Choerocampina, in genera including Hippotion, Hyles and Theretra where labial segment 2 is swollen, hollowed, and its inner surface largely devoid of scaling (Roeder 1972; Göpfert and Wasserthal 1999a; Göpfert et al. 2002), and is an attribute defining the subtribe (Kawahara and Barber 2015). A similar system is found in all Acherontiina (Acherontiini) in genera such as Acherontia, Coelonia and Agrius, and in Xanthopan (Cocytiina, Sphingini), where the inner surface of segment 2 has a depression with modified scaling for receiving sound instead of a tympanum-like plate, and the pilifer is basally hinged rather than fixed (Göpfert and Wasserthal 1999a, b; Göpfert et al. 2002). Both systems are sensitive only to ultrasound and neither is directional; indeed if used for detecting bats they would only report the presence of bats. Minet and Surlykke (2003) provide a detailed overview of hearing in moths including hawkmoths. Thorax (Fig. 5) The thorax has a complex array of sclerites and internal muscles to facilitate movement of the wings and legs. The prothorax carries the anterior legs and is by far the smallest of the three thoracic segments. Dorsally on the prothorax are the patagia, a pair of articulated plates often diagnostically important in Lepidoptera but quite small and of limited significance in hawkmoths. The mesothorax on the other hand is very large as it supports the large forewings chiefly responsible for flight and the midlegs. Dorsally, the large mesothorax comprises two sclerites, the dominating mesoscutum and the posterior mesoscutellum. Covering much of the mesoscutum are the two tegulae, shield-shaped flaps that cover and protect the more delicate flexible areas allowing wing movement. Similarly the metathorax dorsally comprises two sclerites, the metascutum, split along the dorsal midline, and the metascutellum. The sclerites of the lower half of the thorax are far more complex and mainly associated with the legs. There are two thoracic spiracles, one on the membrane between the prothorax and mesothorax and another on the membrane between the wing bases of the mesothorax and metathorax, the latter concealed deeply between the two segments. A detailed account of thoracic sclerites and musculature can be found in Kristensen (2003a). Wings (Figs 11–15) Wing patterns are important diagnostic features at species level in Lepidoptera and the position of markings and bands is described in relation to their distance from the wing base, wing apex or wing margins (Fig. 12). Forewing banding tends to be either transverse between the costal margin and inner margin (Fig 12), or between the apex and inner margin. In both there is a tendency for symmetry in the number of bands about the centre of the wing. The most important bands are generally referred to as (from wing base to apex) the basal, antemedial, medial, postmedial, submarginal and marginal bands (Fig. 12). Spots are uncommon and when present are usually basal or as a discal spot on the forewing or in the vicinity of the tornus in the hindwing, the latter often illdefined. Nijhout (2003) provides a detailed account of the expression of lepidopteran wing patterns. The forewing tends to be long and narrow and the hindwing is always much smaller. Both are fully scaled in most species but in Cephonodes and many Hemaris the majority of scales are shed soon after emergence leaving the wings mostly hyaline. Yoshida et al. (1997) and Yoshida (2005) found that the hyaline wing surface in C. hylas was covered in an array of 190613 Hawkmoths of Australia 3pp.indd 7 7 highly ordered nano-sized protuberances that substantially reduced reflection making the wings difficult to see. In a few genera, including Eupanacra and Cizara, the forewings have a small ‘window’ of translucent white scales. In most species, there is a patch of modified scales that are smaller and smoother on the hindwing upperside (Fig. 12, as ‘low friction scales’) and forewing underside where the wings overlap during flight, and which may help the wings move smoothly over each other. Wing shape varies somewhat between tribes and genera, mainly in the forewing apex which may be falcate to varying degrees, in the extension of the anal lobe and in crenulation of the outer margin. In the forewing the radius R and the radial sector veins Rs1, Rs2 and Rs3 run closely parallel to each other, and the base of the median vein M2 is always closer to the base of M3 than to that of M1 (Fig. 11). Anal veins 1A and 2A fuse to a single vein beyond their bases in the forewing and the posterior cubital vein CuP is absent. The forewing discal cell does not reach midlength of the wing, and its distal margin (formed by the discocellular cross veins) is straight or gently curved. In the hindwing R crosses to the subcostal vein Sc near midlength of the cell, and M1 to CuA2 each arise separately from the cell (Fig. 11). At the apex of the entirely fused 1A+2A in the hindwing the margin protrudes a little forming the tornus or anal angle, and as in the forewing CuP is missing. The hindwing discal cell is always short. The frenulum in males is a single strong bristle arising ventrally from the base of the hindwing (Fig. 13) and is present in all species although vestigial in some Smerinthini including the Australian endemic Coequosa triangularis. It engages with the hook-like, membranous retinaculum behind the forewing costa and joins the wings during flight. In females, the frenulum is a cluster of fine bristles atop a short rod-like pedestal of varying length (long in Gnathothlibus) that engage with a retinaculum consisting of a dense row of soft hairs along the anterior cubital vein CuA (Fig. 14). The retinaculum in females is variable in its development and is sometimes substantially reduced suggesting it may not be effective in holding the frenulum. In many species a tuft of more substantial hairs radiates from near the base of CuA and covers the frenulum and retinaculum, and may have a function associated with those structures (Fig. 15). Legs (Figs 5–9) In basic structure the legs are similar to those of other Lepidoptera and have five segments, the coxa, trochanter, femur, tibia and tarsus. The basal coxa in the foreleg articulates with the thorax, but in the mid and hindlegs, where the coxa is clearly divided into an anterior eucoxa and a posterior meron (Fig. 5), it is more or less firmly attached to the thorax. In Gnathothlibus and some allied genera the foreleg coxa has a tuft of long hair-like scales assumed to have a pheromonal function. The trochanter is always very small and acts somewhat like a knee joint between the coxa and the femur, although in Manduca it is partly fused with the femur (Eaton 1988). The femur is always long and carries no spines of special note. The tibia in the foreleg has an epiphysis (Fig. 6), an articulated flap-like appendage used for cleaning the antennae and proboscis by drawing them through the gap between the tibia and epiphysis which is lined with setae for this purpose. The foretibia in Chelacnema, Hopliocnema, Cephonodes cunninghami and some other species also has a strong apical projection, claw-like in Chelacnema and Hopliocnema but straight and pointed in C. cunninghami. This structure, referred to as a thorn by Rothschild and Jordan (1903), is found in many lepidopteran families, mostly in species associated 29/08/19 11:11:00.75 8 HAWKMOTHS OF AUSTRALIA Figs 2–10. Adult. (2) Head, Cizara ardeniae, lateral. (3) Left labial palp, Psilogramma menephron nebulosa, inner surface. (4) Head, P. m. nebulosa, ventral. (5) Thorax, P. m. nebulosa, lateral with scales removed (c–coxa, em–epimeron). (6–8) Legs, Chelacnema ochra, lateral. (9) Pretarsal claws, P. m. nebulosa, ventral. (10) Abdomen, P. m. nebulosa, lateral with scales removed. 190613 Hawkmoths of Australia 3pp.indd 8 29/08/19 11:11:01.52 STRUCTURE AND FUNCTION 9 Figs 11–15. Wings. (11) Venation, fore- and hindwings, Theretra latreillii, scales removed. (12) Markings, fore- and hindwings, diagrammatic. (13) Male frenulum and retinaculum, G. eras. (14) Female frenulum and retinaculum, Gnathothlibus eras. (15) Female hair tuft covering frenulum and retinaculum, G. eras. A–anal vein, C–costa, Cu–cubital vein, CuA anterior cubital vein, CuP–posterior cubital vein, D–discocellular vein, Fr–frenulum, M–median vein, R–radius, Rs–radial sector vein, Sc–subcosta. 190613 Hawkmoths of Australia 3pp.indd 9 29/08/19 11:11:02.28 10 HAWKMOTHS OF AUSTRALIA with arid environments, and may be an adaptation for assisting emergence from compacted soil. The midtibia carries a pair of apical spurs and the hindtibia an apical and medial pair, but the latter is sometimes missing. Spurs differ from spines in that they are larger, bear sensilla, and have basal articulation. The tarsus has five sections called tarsomeres, the basal one being the basitarsus, is always the longest. Although the tarsomeres look like segments, they are not segments in the true sense as there is only one tarsal muscle. The basitarsus of the midleg often bears a midtarsal comb, a row of long setae along the inner basal half of the segment. Attached at the end of the legs are a pair of independently articulated claws known as pretarsal claws which, together with associated structures form the pretarsus. In many genera there is a sensory arolium (the pulvillus of Rothschild and Jordan 1903) (Fig. 9), a mostly membranous, peg-like structure located basally between the claws. Flanking the arolium is a pair of pulvilli (the paronychia of Rothschild and Jordan 1903) (Fig. 9) that may be single, bilobed or absent. Rothschild and Jordan (1903) and Kristensen (2003a) provide very detailed accounts of leg morphology. Abdomen (Fig. 10) In the male the first eight segments form the abdomen proper while segments 9 and 10 comprise the genital structures. In the female only the first seven segments are of ordinary appearance, segments 8–10 are modified for the reproductive system. In the male segments 1–8 each have their tergite and sternite separated by a defined pleuron that is membranous except in segment 1. Segment 1 is small and the sternite ill-defined, vestigial or absent. The first abdominal spiracle lies on the intersegmental membrane immediately anterior of the pleuron on segment 1, while the spiracle of segment 2 is located on the tergite, that on segment 3 partially on the tergite and those on segments 4–7 on the pleuron. There is no spiracle on segment 8. The structure of the first and last sternites shows notable modification in some genera. In the most anterior sternite (actually sternite II, as I is never identifiable) the shape is often variable between genera while the last sternite (sternite VIII in males, sternite VII in females) is always without spines and can vary considerably in shape between some genera. Much of this variation is discussed by Rothschild and Jordan (1903). Tergite 8 in males may be either evenly sclerotised or divided into a medial and two lateral sclerites joined by membrane. The hair-like scales attached to a tripartite tergite are thus divided into three clusters forming a fantail as in Macroglossum. Notable in the subfamilies Sphinginae and Macroglossinae are small spine-like scales bordering the posterior margin of the segments, on tergites 2–8 in males and 2–7 in females, and on sternites II–VII. Those bordering the tergites are usually denser and stronger than those on the sternites, and may be arranged in single or multiple overlapping rows. Rothschild and Jordan (1903) identified three kinds of spination and discuss these in some detail. In the male of most hawkmoths there is a pair of andronical tufts or brushes (hairpencils) (missing in many Smerinthinae, e.g. Cypa, Kitching and Cadiou 2000). These are situated on the anterolateral corner of sternite III and at rest lie within a slitlike pocket across sternites II and III. They are everted to emit pheromones, possibly for attracting females or during courtship, but their function has never been confirmed. Hawkmoths lack abdominal tympanal organs as found in many other moth families. Male external genitalia (Figs 16–21) The external male genitalia comprise mostly heavily sclerotised structures that are primarily modifications derived from 190613 Hawkmoths of Australia 3pp.indd 10 abdominal segments 9 and 10. These structures often provide important diagnostic characters for distinguishing species and genera, and sometimes tribes. The frame of the external genitalia consists of the tegumen, which is derived from tergite 9, and the vinculum, which is derived from sternite 9, and, as upper and lower halves, they form a sclerotised, transverse ring to which other genital structures attach (Figs 17, 20). The vinculum is attached to the tegumen by an extension that lies along the anterior edge of the tegumen and forms a flexible connection. The base of the vinculum is extended anteriorly and medially to form the saccus, a protrusion carrying muscle attachments for movement of the copulatory phallus (Fig. 17). The uncus and gnathos attach to the distal part of the tegumen and together often appear as a beak-like structure (Fig. 17). Emerging between them is the short membranous anal tube, the end of the digestive system. In most hawkmoths, the uncus is undivided but in some it is partially or completely bifid, as in all Psilogramma species. Usually the uncus tapers to a bluntly pointed apex that is often downturned but sometimes it is more elaborate and laterally expanded. The gnathos is mostly well developed although it is never as long as the uncus. Usually it is rounded ventrally and flattish dorsally, and sometimes apically pointed or bifurcate, as in Ambulyx, Amplypterus and Cerberonoton. In a few species it is substantially reduced as in Acherontia and some Megacorma and Pseudoangonyx, and occasionally so much so as to be effectively absent, as in Cephonodes. The paired valvae, articulating with the vinculum, are the clasping organs used for grasping the female during copulation. They are large flat structures, generally ovate in shape and when at rest enfold the uncus, gnathos and phallus. Basally along the ventral margin of the valva is the swollen sacculus that usually is extended distally as the harpe (Figs 16–19). The harpe is usually well-developed, often lanceolate and variously spined, and frequently diagnostic at species level. Rarely is it absent, as in Psilogramma. Each valva is manipulated by muscles attached to an apodeme, often a wellsclerotised process (Figs 16, 18, 19). This structure, sometimes wrongly interpreted as a transtilla in the Sphingidae, is homologous with what is present in most other Bombycoidea (Zwick 2009), and we follow Zwick in calling it an apodeme. The valvae of some genera carry additional structures such as lobes, spines and pouches, particularly well developed in genera such as Cypa and Tetrachroa (Fig. 18). A low medial pouch-like structure termed an ampulla (Fig. 19) is frequently encountered. In a few genera the genitalia are asymmetrical to varying degrees, especially noticeable in the valvae of Cephonodes (Fig. 19) which, as in some other genera with asymmetrical genitalia, also have the uncus twisted to the left and the gnathos to the right. There is a corresponding asymmetry in the females of those genera, but in the opposite direction. In many species the outer surface of the valva bears a patch of friction scales (Figs 16, 17), which act as a stridulatory organ. In the Sphingini these are rubbed against needle-like spines on the posterior edge of the eighth tergite, in the Ambulycini against a curtain of scales laterally on tergite 8 and in other tribes they act in other ways, some yet to be documented (I.J. Kitching pers. comm.). Although the function of this sound production is not entirely understood, one purpose appears to be to jam the ‘radar’ of predatory bats, but it may also be used to communicate during mating (Mell 1922; Lloyd 1974; Nässig and Lüttgen 1988; Nässig et al. 1992; Kitching and Cadiou 2000; Kawahara and Barber 2015). The friction scales can be large and numerous as in Agrius and 29/08/19 11:11:02.35 STRUCTURE AND FUNCTION 11 Figs 16–21. Male genitalia. (16) Ventral, Hippotion johanna. (17) Lateral with left valva removed, Hippotion johanna. (18) Ventral, Tetrachroa edwardsi. (19) Ventral, Cephonodes australis. (20) Lateral with left valva removed showing membranous manica and anellus, Psilogramma argos. (21) Phallus with vesica everted, Macroglossum alcedo. Psilogramma, but may be microscopic if present in Smerinthinae. In the Choerocampina (e.g. Hippotion, Figs 16– 17) and some Macroglossinae (e.g. Acosmeryx and Daphnis) they are large but few in number. In some genera with large friction scales (e.g. in Cerberonoton and Psilogramma) the sound can be clearly audible (Robinson and Robinson 1972; 190613 Hawkmoths of Australia 3pp.indd 11 Nässig and Lüttgen 1988; Nässig et al. 1992; Kitching and Cadiou 2000), but in many others it is ultrasonic. Friction scales are absent in some Smerinthinae, all New World Sphinginae, many small species (e.g. small Macroglossum, Sphingonaepiopsis, Neogurelca) and diurnal species (e.g. Cephonodes and Hemaris). 29/08/19 11:11:03.05 12 HAWKMOTHS OF AUSTRALIA The membranous diaphragm closes the body cavity between the valva bases, with some regions sclerotised and carrying muscle attachments. The juxta is a small sclerotised structure below the phallus that serves as a ventral support for the phallus and has retractor muscles attached. It is present in all the Sphingidae but in many Macroglossinae in particular it is quite small, often a lightly sclerotised triangular or rhomboidal sclerite. The juxta is larger in Sphinginae and very obvious in most Smerinthinae. The phallus, the organ of copulation, is also widely known as the aedeagus, a term considered inappropriate for use in the Lepidoptera (Kristensen 2003b) as the aedeagus of other orders is believed not to be homologous. The phallus originates from the bottom of a pocket-shaped invagination in the diaphragm and is supported in part from below by the juxta. The surrounding diaphragm is modified to form the sleevelike inner manica and the surrounding anellus encapsulating the phallus and allowing it to slide back and forth (Fig. 20). The manica attaches as a ring around the phallus, basal to its midlength. Basally the phallus is excavated across its dorsal half, leaving the lower half known as the coecum for muscle attachments for manipulating the phallus. The ductus ejaculatorius, the tube carrying sperm from the testes, passes through the diaphragm and enters the phallus above the coecum and continues through the inside of the phallus to its apex, forming the eversible vesica. The vesica, usually thin and membranous (Fig. 21), is everted during copulation and withdrawn within the phallus when at rest whereupon the ductus ejaculatorius is compressed by contracting to the base of the phallus. The opening at the apex of the phallus when the vesica is withdrawn creates a false gonopore or secondary gonopore. The vesica may have up to three diverticula and often bears sclerotised spines or rods termed cornuti, often species specific. Apically the phallus is usually ornamented with spines of varying form which are also often species specific. In some Theretra and some Cechenena they are in the shape of a longitudinal row of fine crisscrossed spines in the distal third (Pls 91, figs c-h; 92, figs b, d; best seen in 91, g, d and 92 b). These are deciduous and can be left behind in the female after mating (Rothschild and Jordan 1903: lxxxii), their purpose perhaps being to prevent subsequent matings, although this is unconfirmed. Male internal reproductive system (Figs 22–23) The internal reproductive system appears as a tangled mass of ducts of remarkable length occupying much of abdominal segments 4−8. Attached to the phallus is the ductus ejaculatorius, the longest of all the ducts that can be as much as five times the length of the abdomen. The posterior portion is always straight and positioned dorsally in the abdomen, and houses the spermatophore (a capsule containing sperm), which is transferred to the female during copulation. At its other end it branches twice forming the paired vas deferentia, each of which leads to the paired but externally fused testes. Situated part way along each vas deferens is a small swelling, the seminal vesicle, a temporary store for the descending spermatozoa. Each vas deferens is paired at its base with an accessory gland. Female reproductive system (Figs 24–27) As with all but the most primitive moths, sphingid female genitalia are ditrysian, i.e., they have separate openings for copulation and oviposition. The copulatory opening, the ostium bursae, lies ventrally on segment 8 (Fig. 27). During Figs 22–23. Male reproductive system, Psilogramma menephron nebulosa. (22) In situ, dorsal view with tergites and extraneous tissues removed. (23) Expanded. 190613 Hawkmoths of Australia 3pp.indd 12 29/08/19 11:11:03.41 STRUCTURE AND FUNCTION 13 Figs 24–27. Female reproductive system, Psilogramma menephron nebulosa. (24) Female, expanded, entire, dorsal view. (25) Female, expanded, partial, lateral view. (26) Female terminalia and bursa copulatrix, dorsal view. (27) Female terminalia, ventral view. Images 24, 26 and 27 are orientated with terminalia at the top following traditional convention. copulation males transfer a spermatophore into the female’s large sack-like bursa copulatrix, differentiated into a narrow proximal ductus bursae and a distal corpus bursae which often carries a signum, a scobinate patch or bands of very small spine-like tubercles that protrude on the inner wall of the corpus bursa and is often diagnostic for species or genera (Fig. 26). The function of the signum is to rupture the external wall of the spermatophore that is subsequently digested (Galicia et al. 2008), a process not required for fertilisation as spermatozoa are released before rupture of the spermatophore (Drummond 1984). Spermatozoa leave the bursa copulatrix and pass through the ductus seminalis into the bulbous spermatheca located near the base of the spermathecal gland, where they are stored. The ductus seminalis normally attaches to the ductus bursae dorsally and leads to the common oviduct where it joins laterally adjacent to the spermathecal duct that attaches dorsally to the common oviduct (Fig. 25). Within the ductus bursae and adjacent to the ostium bursae are the antrum and colliculum (neither figured). The antrum may not be anatomically part of the ductus bursae but rather an invagination of the sterigma. The colliculum, a sphincterlike structure at the anterior end of the antrum probably serves to prevent spermatozoa from being inadvertently expelled through the ostium bursae after their release from the spermatophore. It would seem logical for the ductus seminalis to go directly upward from the ductus bursae to the common oviduct that sits above it, as in Gnathothlibus, but it appears 190613 Hawkmoths of Australia 3pp.indd 13 that in many species the antrum, and consequently the ductus bursae, have rotated causing the ductus seminalis to first loop around the ductus bursae. Kitching (2002) first discovered this rotation which he found across a wide range of species to varying degrees but always in a clockwise direction. The function of the twisting is unknown but can be extreme, sometimes more than one and a half turns. In Psilogramma menephron, for example, the ductus bursae has rotated through 360º so that the ductus seminalis turns down along the right-hand side of the ductus bursae, passes under it, then up its left side and then up to the common oviduct (Fig. 25). Eggs are produced in the paired ovaries (Fig. 24). Each ovary has four ovarioles, their basal dividing branches forming the calyx. Oocytes (developing eggs) form in the ovarioles before passing down the common oviduct at maturity. As the eggs move down the common oviduct they are fertilised by the spermatozoa stored in the spermatheca and as they continue further past the shared duct of the paired accessory glands (also attached dorsally to the common oviduct) they receive a secretion that glues the eggs to the substrate. Eggs are carefully placed with the aid of the paired, bulbous papillae anales that carry sensory setae for touch and smell and which flank the ovipore that lies immediately below the anus (Fig. 27). Each papilla analis is extended basally into a long, rod-like posterior apophysis to which muscles attach apically that are used to manipulate and retract the ovipositor during oviposition. Extension of the ovipositor is probably by haemostatic pressure. The sclerotised plates surrounding the 29/08/19 11:11:03.86 14 HAWKMOTHS OF AUSTRALIA ostium bursae are derived from sternite VIII and together form the sterigma, the portion anterior to the ostium bursae termed the lamella antevaginalis and the posterior one the lamella postvaginalis (Fig. 27). Tergite 8 lies above the ostium bursae (Fig. 26) but in situ it is hidden beneath tergite 7. The long, rod-like anterior apophyses are forked basally, one branch fused to the anterolateral corner of the lamella postvaginalis and the other to the anterolateral corner of tergite 8. They serve to manipulate these sclerites abutting the copulatory opening during copulation by muscles attached apically (Kristensen 2003a, after Kuznetzov and Stekolnikov 2001). The female reproductive system provides comparatively few species diagnostic characters in contrast to the male genitalia. They are more useful at family and subfamily level. In hawkmoths Kitching and Cadiou (2000) found that the female genitalia generally conform to one of two types. In Sphingulini and many Smerinthini the ductus bursae is short and thick-walled while the corpus bursae is small and without a signum and there is a distinct ‘neck’ between the ductus bursae and the corpus bursae. This type of female genitalia is confined within abdominal segments 6 and 7. The other type, described by Kitching and Cadiou, is found in other Smerinthini, Ambulycini, Sphinginae and Macroglossinae (Figs 24, 26). The ductus bursae is long and membranous with no defined junction between the ductus bursae and corpus bursae which is an elongated oval sac usually with a signum. Such genitalia extend almost the full length of the abdomen. Because the majority of structures are unsclerotised, care needs to be taken in assessing characters that appear different in dried or fresh specimens, and especially between mated and unmated females. The most useful structure for separating species is the sclerotised signum on the corpus bursae, usually a single or double band of very small, sclerotised, conical tubercles, but sometimes a small patch or entirely absent (Fig. 26). Egg Eggs of hawkmoths are usually subspherical, slightly flattened dorsally and ventrally, but some are slightly elongated such as those of the Australian endemic genera Hopliocnema, Chelacnema and Synoecha. The outer egg shell, the chorion, is nearly smooth but with a fine net-like sculpturing in a rosettelike pattern radiating from the inconspicuous micropyle (Fig. 28). This pattern results from the imprints of the follicular cells that deposited the chorion (Orfanidou et al. 1992). The axis of the egg is horizontal, and the micropyle is lateral. The micropyle allows entry of sperm cells for fertilization, and the micropyle has to be orientated towards the spermathecal duct during oviposition (Fehrenbach 2003). The surface has aeropyles, minute holes to allow air to flow to a thin air layer below the chorion (Fig. 29). Oxygen intake for the embryo is by diffusion, and the complex process between water balance and oxygen intake is detailed by Fehrenbach (2003) and Woods et al. (2005). In many hawkmoth eggs the surface also bears low rounded protuberances of unknown purpose, usually situated in a shallow depression, either scattered or in clusters (Fig. 29). Surface structures of the chorion of hawkmoths have been illustrated and discussed by Danner et al. (1998) based on scanning electron micrographs for species in 17 Palearctic genera, in an attempt to use these structures to distinguish species. While it is possible to identify the eggs of many lepidopteran species from features of the chorion, these characters are subtle in hawkmoths, and Kitching and Cadiou (2000: 117, note 264) advise caution in the interpretation of perceived differences in these structures. The chorions of 190613 Hawkmoths of Australia 3pp.indd 14 additional species of Daphnis, Gnathothlibus and Megacorma are figured by Eitschberger (2008a, 2008b, 2008c). Larva Hasenfuss and Kristensen (2003) provided a comprehensive account of the larval lepidopteran skeleton and musculature. The larval lepidopteran digestive and excretory systems are detailed by Dauberschmidt (1933) and Barbehenn and Kristensen (2003), including figures of Sphinx and Daphnis. Wasserthal (2003) described the respiratory system in lepidopteran larvae, and Yack and Homberg (2003) the nervous system. Peterson (1912) gave detailed accounts and figures of the respiratory, nervous and digestive systems of Manduca sexta. Head (Figs 34, 35) The larval head is always rounded in the 1st instar but in some species may be exaggeratedly elongated dorsally from the second to the penultimate instar, as in Coequosa. In those species the head is usually less elongate and somewhat rounded at the vertex in the last instar. In its dorsal/posterior half the head capsule or epicranium is bisected longitudinally by the medial adfrontal suture (or coronal suture), which forks anteriorly as the lateral adfrontal sutures to enclose the frontoclypeus (Fig. 34). The epicranium is strengthened internally below the adfrontal sutures by an inverted Y-shaped ridge that provides support for the muscles operating the mouthparts. In many moths the head capsule in the last instar splits along lines of weakness adjacent to the adfrontal sutures at ecdysis but in hawkmoths it is shed intact. On the lower lateral surface on each side of the epicranium are six stemmata (incorrectly called ocelli), which are single lens visual organs. The most prominent mouthparts are the chewing mandibles, which are always darkened apically by heavy sclerotisation. They are protected above by the labrum, which joins and articulates with the anteclypeus. Below the mandibles are the sensory maxillary palps, which assist in evaluating food before consumption. Unlike the maxillary palps, the labial palps are very small and lie between the bases of the maxillary palps, flanking the similarly small spinneret. The spinneret, used for extruding silk from modified salivary glands in all instars, is most obviously used by those last instar larvae that construct loose silken cocoons. Flanking the mouthparts are the sensory 3-segmented antennae (Fig. 35). The head carries many sensory setae, developed in all instars but longest in the 1st instar. Thorax and abdomen (Figs 30–33) The three thoracic segments (prothorax, mesothorax and metathorax) and 10 abdominal segments are clearly defined in all instars (Fig. 31). Spiracles are present on the prothorax and on abdominal segments 1−8. The body carries many setae arising from small tubercles. The setae are most developed in the 1st instar and although they increase in number in subsequent instars they decrease in size so that by the last instar in most species few if any are detectable. These setae are important in lepidopteran classification, particularly at the family rank. They are usually defined in a standard terminology primarily by their position and portrayed on setal maps; their study known as chaetotaxy. Common (1990) provides an easy to follow overview of moth chaetotaxy, and Hinton (1946) summarises the chaetotaxy of hawkmoth larvae and its differences from other Lepidoptera families. In hawkmoths the positions of the primary setae are always the same (Fig. 32) but their size and shape can vary between genera and sometimes between species. Such differences are most obvious in the 1st instar. The setae are usually apically 29/08/19 11:11:03.93 STRUCTURE AND FUNCTION 15 Figs 28–29. Scanning electron micrographs of the chorion, Daphnis dohertyi. (a) Lateral micropyle showing the radiating, rosette-like, fine sculpturing typical of many sphingid eggs. (b) Enlargement of the surface showing the aeropyles and clusters of low rounded protuberances. Images Ulf Eitschberger. bifurcate to varying degrees, with the most extreme examples found in Hemaris and Cephonodes (Fig. 32). As larvae pass through their successive instars the low tubercles bearing the setae usually diminish in size with a few exceptions where they remain enlarged, mostly on the dorsal thorax, as in Psilogramma. In exceptional cases they actually increase in size to take on the appearance of spines, as in Coequosa. The development of these tubercles is often diagnostic at the species level. In most genera where the tubercles diminish in size their position remains highlighted as a pale spot that gives the larva a somewhat mottled appearance. Each thoracic segment carries a pair of jointed true legs (Fig. 33), the equivalent of the true legs in adults, which are used by older larvae to hold the leaf edge while feeding. The prothorax bears a sclerotised dorsal plate, the prothoracic shield, which in all instars always has transverse rows of sometimes quite large tubercles (Fig. 31), each of which bears a single seta. Abdominal segments 3−6 and segment 10 each carry a pair of unsegmented prolegs, those on segments 3−6 are known as ventral prolegs and those on segment 10 as claspers or anal prolegs (Fig. 31). The ventral prolegs are mainly used in walking and the claspers, as their name implies, for grasping. In the 1st instar the ventral prolegs on segment 6 are always larger than those on segments 3−5 but this gradually becomes less pronounced in subsequent instars until by the penultimate there is no discernible difference. All prolegs have a sclerotised plate laterally on their outer surface, the lateral shield, which provides attachment internally for muscles assisting movement of the proleg. Apically on the prolegs is a planta, a sole-like pad that carries many small hooked spines or crochets used for gripping. In hawkmoths these are arranged in two transverse rows although sometimes the inner row is much reduced. Apart from the claspers, segment 10 also has a sclerotised anal plate that protects the anus below. The anal plate always bears raised tubercles, the two medial ones of which are often the largest on the larva and the apex of the anal plate may be either rounded or bi-lobed. Segment 8 bears the caudal horn that is often useful in distinguishing species (Figs 30–32). It takes many forms and may be slender or robust, curved forwards or backwards, or straight, and ranges from very long to very short or even absent. In later instars the caudal horn often sits atop a tumidity, a fleshy dorsal elongation of abdominal segment 8, well developed in diverse genera such as Psilogramma, Gnathothlibus, Macroglossum, Cephonodes and some Theretra species (notably T. silhetensis). Rarely the caudal horn is absent from the 1st instar, as in Coequosa australasiae. In two other 190613 Hawkmoths of Australia 3pp.indd 15 Australian endemics, Coequosa triangularis and Hopliocnema brachycera, it diminishes in size through the instars to being absent in the last one. It is always forked apically in the 1st instar as a result of two tubercles meeting basally in a V-shape, but in successive instars these tubercles are reduced in size diminishing the forked effect. In all instars, and along its entire length, the caudal horn also carries many short tubercles that are often spine-like and, like those at the apex, each bears a short terminal seta. Usually the horn is held erect but in the early instars of many Macroglossum species and some others it is held horizontal. Last instar Theretra oldenlandiae are well known for waving their white-tipped horn back and forth as they move along. Damage to the caudal horn seems not to affect the development of the larva. However, the actual purpose, if any, of the caudal horn in sphingids remains unexplained. It may, in fact, be the last survivor of a larger compliment of scoli, as seen in other Bombycoidea, viz. Saturniidae and Brahmaeidae (I.J. Kitching pers. comm.). It is structurally homologous with scoli, a cuticular protuberance that itself bears tubercles which bear setae. In the Macroglossinae, except in Cephonodes and closely allied genera and in the tribe Dilophonotini, the head is small and the anterior segments are retractable. In some genera, including Agrius and Psilogramma, the abdominal segments have multiple transverse folds but in most others the abdominal segments are largely or completely smooth. Among Australian genera the metathorax and abdominal segment 1 are sublaterally expanded into a fleshy ridge in Acosmeryx, and the eyespot on abdominal segment 1 in Theretra queenslandi is considerably swollen. Eyespots, if present, are always positioned against the anterior margin of a segment (Fig. 30). Pupa Pupae tend to be fusiform although most are bluntly rounded at the head and pointed at the rear. The head is dominated by the proboscis that usually continues as a narrow linear sheath between the wings to their apices but never beyond (Fig. 37). In a few genera the proboscis is very short and terminates before or on reaching the wings. In many Macroglossinae, the anterior portion of the proboscis is keel-like and may project in front and below the head (Fig. 38). However, in the Sphinginae, and derived independently in the genus Pergesa (Macroglossinae), the proboscis has a trunk-like extension that is sometimes recurved (Fig. 39). With the development of the adult moth within the pupal skin, the adult proboscis is looped back upon itself within the proboscis appendage 29/08/19 11:11:04.02 16 HAWKMOTHS OF AUSTRALIA Figs 30–35. Larva. (30) General head and body markings. (31) External morphology. (32) Chaetotaxy of 1st instar (Cephonodes kingii) setal groups: D-dorsal, L-lateral, SV-subventral, XD-tactile dorsal; numbers distinguish individual setae belonging to the same group. (33) True leg. (34) Head, anterior. (35) Head, lateral. and thereafter continues to the apex of the pupal proboscis ventrally along the body (Fig. 40). As the larva transforms to a pupa this trunk-like extension begins very small and tightly coiled but quickly expands over an hour or two to reach its 190613 Hawkmoths of Australia 3pp.indd 16 full length (Pl. 14, figs g–l). The antennae and legs are fused to each other and to the body but the hindlegs are hidden (Fig. 37). The thorax is visible only dorsally except for very small sections of the prosternum and mesosternum that are 29/08/19 11:11:05.80 STRUCTURE AND FUNCTION 17 Figs 36−43. Pupa. (36) Dorsal (Theretra latreillii). (37) Ventral (Theretra latreillii). (38) Lateral (Theretra latreillii). (39) Lateral (Psilogramma casuarinae). (40) Lateral, showing location of the developed adult proboscis within the pupa (Psilogramma casuarinae). (41) Anterior of pupa showing position of the thoracic spiracle, lateral (Psilogramma casuarinae). (42) Male pupa, terminal segments, ventral (Theretra latreillii). (43) Female pupa, terminal segments, ventral (Theretra latreillii). visible ventrally. The prothoracic spiracle is situated at the posterior margin of the prothorax and is partly or entirely obscured by a flat spiracle cover (Fig. 41). On the dorsal metathorax is the metathoracic plate, a raised, sculptured, transverse band often divided at the dorsal midline and developed slightly differently between genera (Fig. 36). The forewings are large and usually reach the posterior margin of abdominal segment 4. Unlike the head, thorax and abdomen they have no sculpturing and are often partly translucent. The forewings mostly conceal the hindwings, which are 190613 Hawkmoths of Australia 3pp.indd 17 visible laterally only as far as abdominal segment 3 and in some genera also as a narrow sliver ventrally against the forewing termen. There are 10 abdominal segments, the last of which forms the spined cremaster used to assist in anchoring the pupa during pupation. In some deep-burrowing subterranean species, the cremaster may also assist as the pupa works its way to the soil surface prior to adult emergence. The spination is often useful in distinguishing species (compare Figs 53−64, p. 42). The cremaster and its base originate from the larval 29/08/19 11:11:06.61 18 HAWKMOTHS OF AUSTRALIA anal plate, and ventrally on either side are the rounded remnants of the claspers bordering the anal cavity, which is often deeply recessed (Figs 42−43). Anterior of the cremaster on the ventral surface are genital ‘scars’, the developing genital opening only on segment 9 in the male (Fig. 42), on both segments 8 and 9 in the female, the developing ovipositional and genital openings respectively (Fig. 43). The intersegmental membrane adjacent to abdominal segments 5, 6 and 7 allows movement of those segments, necessary during pupation and emergence of the adult. Laterally, and anterior 190613 Hawkmoths of Australia 3pp.indd 18 of the spiracle on segments 5 and 6, and sometimes on 7, are defined grooves or sculptured ridges, the spiracular furrows of Kitching (2002). Oberprieler and Duke (1994) suggested that these spiracular furrows assist subterranean pupae in moving to the surface prior to adult emergence. Spiracles are present on abdominal segments 1−8 although that on segment 1 is nearly always hidden below the edge of the forewing. The larval caudal horn is usually marked by a scar on segment 8, and sometimes the prolegs of abdominal segments 5 and 6 show similar scars. 29/08/19 11:11:06.63 Collection and preservation The study of hawkmoths can be a fascinating and rewarding pursuit and establishing a collection can contribute significantly to our knowledge of systematics and zoogeography. The majority of museum collections of hawkmoths have originated from dedicated and enthusiastic amateurs who have donated their collections. However, to be scientifically useful a collection must be properly presented and curated. The following notes are designed to give guidance on collecting, preparing and maintaining a collection. We do not address all aspects of collecting and preserving hawkmoths where such information relates to moth collecting in general and the information is readily available elsewhere, e.g. setting, relaxing, labelling and maintaining a collection. Excellent texts in this regard are Upton and Mantle (2010) and May (2014). Here we concentrate on techniques directly relevant to hawkmoth collecting and study. Collecting adult hawkmoths Netting. Although often neglected, collecting from flowers during late afternoon and at dusk can be rewarding for sphingids not often otherwise encountered, such as species of Macroglossum. Even day-flying Cephonodes species can be taken at dusk at flowers. Favoured flowers include Lantana, purple snakeweed, Pentas, pawpaw, duranta and abelia. Light trapping. Hawkmoths, like many other insects, can use the ultraviolet light spectrum. The ultraviolet light from some mercury vapour lamps will readily attract hawkmoths at night. Clear lamps, which lack the white coating designed to reduce ultraviolet emission, are the most efficient in attracting insects. While pure mercury vapour lamps require a choke (also known as a ballast), blended mercury vapour lamps need no auxiliary equipment. Both kinds of mercury vapour lamps require a 240 volt power supply. The advantage of pure mercury vapour lamps is that they are available in much higher wattages with the potential to attract more moths. While pure mercury vapour lamps are available in wattages ranging from 125–2000 watts, 250 watt lamps are perhaps the most practical, given that higher wattages do not proportionally attract more insects. The ultraviolet light produced by these lamps is harmful to human eyes, and it is essential to wear glasses with lenses that filter out ultraviolet radiation. A gritty feeling in the eyes is a symptom of ultraviolet damage, and exposure can pose a serious risk. However, the demise of mercury vapour lamps may be pending with the advent of LED lighting and the above-mentioned lamps are already difficult to find. Black light or actinic blue fluorescent tubes are a less effective alternative, one of the better ones being the ‘Gecko’ 50 watt folded blacklight tube. Blacklight tubes of 15 or 40 watts also are satisfactory and lower wattages can be operated from either 240 volts or a 12 volt car battery. The disadvantage of black light tubes is that their visible light output is low and they need to be run in conjunction with a white light or torch for the moths’ colours to be seen. To provide a landing site for attracted moths, a white vertical sheet (a bed sheet or shade cloth) should be provided with the light near the top centre so that the moths can land; otherwise, hawkmoths that are attracted to the light may pass by or land in nearby vegetation where they can quickly damage their wings. A ground sheet of similar material should be 190613 Hawkmoths of Australia 3pp.indd 19 positioned below the light, preferably about 4 m square or larger, and placed centrally beneath the vertical sheet. Many species of hawkmoths will settle on the ground sheet; hence the larger the size the better. Even so, some hawkmoths (and many other insects) will settle on surrounding vegetation, so it is helpful to have a little cleared space around the sheet and disturb the vegetation from time to time. Other species may remain active in flight nearby and have to be netted. To be most effective, light sheets should be positioned at the best available vantage point, such as in a forest clearing, or near a stream bank or track verge. Usually a high vantage point is more successful than a low one, so that a light on a hilltop overlooking nearby tree tops would generally be better than a light at the bottom of the hill. Sheltered situations that avoid strong windy conditions are also far more satisfactory. The light should always be placed on the sheltered side of the sheet. A bright moon can substantially reduce the attraction of moths to a light. Try to avoid the week between a half and full moon. The most productive period for running a light is the two weeks beginning one week after a full moon. Different species of hawkmoths tend to fly at different hours of the night, some having one or more periods of activity when they may be attracted to light, many mostly after midnight. For best results, lights should always be continuously operated from dusk until well after dawn. Killing specimens Specimens intended for collections need to be killed quickly to avoid damage from excessive fluttering. There are three preferred ways of killing hawkmoths: by freezing, by injection, or by using a killing jar. Freezing is usually impractical in the field but often useful in dispatching bred adults without the need to handle the specimens. Usually 20 minutes or more are needed to kill hawkmoths. Injecting a killing agent with a hypodermic syringe inserted between the thoracic and abdominal segments is preferred by some lepidopterists, especially for larger species. Ethanol is often used as it does not damage the DNA if the specimen is required for molecular study. Oxalic acid injected in very small amounts as a saturated aqueous solution is an alternative that also does not degrade DNA, and such specimens do not appear to exhibit rigor mortis. An advantage of this approach is that it kills specimens almost instantaneously. Ethyl acetate can also be used but it dissolves most plastic syringes. Killing jars are by far the most widely used, convenient and effective way of killing hawkmoths in the field. There are many killing agents used including ethyl acetate, potassium cyanide, ammonium carbonate and others. Each has its advantages and individuals often have their preference but we deal here only with ethyl acetate, the most commonly used killing agent for hawkmoths. Ethyl acetate provides a relatively safe and convenient way of killing hawkmoths in the field. Poisoning occurs only if the chemical is swallowed, but contact with eyes and open wounds must be avoided. The disadvantage of this method is the need for recharging as the ethyl acetate evaporates; however, dependent on usage, a well-sealed jar should be effective for up to four hours. Use a wide-mouthed, screw-topped glass or plastic jar, but test plastic jars to ensure 29/08/19 11:11:06.68 20 HAWKMOTHS OF AUSTRALIA the ethyl acetate does not dissolve the plastic in question. Prepare the jar by placing a layer of dry cotton wadding on the base, next place a small wad of cotton wool semi-saturated with ethyl acetate and then cover by a further two centimetres or more of dry cotton wool. Care needs be taken not to use too much ethyl acetate, as contact with liquid ethyl acetate will result in scale damage. Alternatively, a piece of kitchen sponge may be fixed to the inner side of the lid of a jar and saturated with ethyl acetate. Collected moths should be left in the killing jar for at least 30 minutes before being removed. By using multiple jars to initially stun specimens before transferring them to a larger jar, the damage caused by newly caught and highly agitated adults can be avoided. Field storage Depending upon the length of a field trip, storage space and personal preference, specimens can be temporarily protected and stored in the field in several ways. Regardless of the method used, field stored specimens must be labelled in a manner that avoids later confusion. Papering specimens is usually the preferred method of storage. Specimens are best placed individually into glassine envelopes (or glassine or paper triangles), with the collecting data written on each. Triangles made from newspaper or equivalent can help absorb moisture and body fats from specimens prone to ‘greasing’. Store in a deep freezer if available to keep specimens soft for later setting. Otherwise, dry the specimens in their envelopes sufficiently to prevent decay and mould and pack into containers such as plastic takeaway food containers, for transport. A teaspoon or so of ‘Dettol’ (or equivalent) or other fungicide such as thymol crystals or chlorocresol will help prevent mould and deter ants, another serious threat to specimens in the field. Pinning specimens should be done before the moth has dried and hardened. The pin is placed vertically through the centre of the thorax and two pins may be placed on either side of the body or at the base of the forewing costa to prevent the moth from rotating. Pinned specimens are best kept in a storebox or other cork or polyethylene foam lined box treated with fungicide as used for papered specimens. Pinning allows more successful relaxing and setting and better protects appendages, but papering saves space. Labelling specimens If a collection is to have scientific value, it is essential that each specimen carry a label(s) on the pin that, at a minimum, includes the locality (e.g. 10 km E of Mareeba, Qld, 16°59’11”S, 145°31’15”E), date of collection (written as 12 Mar. 2017 or 12.iii.2017, not 12.3.17 or 3.12.17) and name of collector. The locality should be accurate enough to establish where the specimen was taken, ideally with a latitude/longitude derived by GPS. Always include the State and, if needed, also the country. In addition, specimens can be numbered for databasing but never attach only a number referring to data recorded separately in a register or database. Sadly, some very valuable collections have been rendered useless by the loss of their accompanying data. Labels are best made from a thin white acid-free card and data written in permanent ink, laser home printed or commercially printed. If desired, further labels can be attached with valuable additional information, such as habitat or the foodplant of reared specimens. Preparing molecular specimens For most systematic studies of hawkmoths using DNA, the so called ‘barcode’ section of the mitochondrial gene COI is used. It requires fragments of more than 658 base pairs (bp) long. But DNA deteriorates rapidly after death, breaking into ever 190613 Hawkmoths of Australia 3pp.indd 20 shorter fragments. To preserve the DNA for subsequent analysis, samples must not only be dried (dehydrated) quickly but ideally also stored in a way that slows further deterioration. As hawkmoths are large, one or two legs are generally sufficient for DNA extraction and sequencing. Ideally, legs should be removed immediately after death and stored in 95% ethanol at low temperature (change ethanol next day to avoid dilution by body fluids). The alcohol not only dehydrates the sample but also keeps it dehydrated, denatures any enzymes that might break down DNA, and prevents degradation and contamination by microbes, fungi and insects. While storing ethanol-preserved samples at room temperature is satisfactory on a temporary basis, deterioration of the DNA continues although at a slow pace, so storing in a freezer is best, the colder the better. If 95% ethanol is unavailable, propylene glycol or automotive glycol-based antifreeze can be used, but lowconcentration ethanol, methylated spirits and rubbing alcohol (isopropanol) are generally not suitable for DNA preservation. Although inferior to ethanol dehydration and storage, if initially dried quickly, leg samples can be also stored dry in airtight vials, ideally in a freezer. Unmounted specimens should not be relaxed for setting prior to leg sample removal as DNA is significantly destroyed by rehydration. Specimens are best killed either by freezing, cyanide or dropping into 95% ethanol. Caution is needed in using ethyl acetate or ammonia-based killing agents as they damage DNA. Label data on both the adult and associated leg samples should be complete and unambiguously linked. Dissecting genitalia Although many species can be identified from colour images of set adults, it is sometimes necessary to dissect and examine the genitalia for distinguishing features, especially when undescribed species are suspected. Usually examination of the male genitalia is sufficient but female dissections can sometimes be productive. The technique of dissection requires practice and one should not be discouraged from unsuccessful early attempts. Male genitalia are best processed as follows: 1) With scissors remove the last third of the abdomen and place in aqueous potassium hydroxide solution (10% KOH), leaving at room temperature for around six hours for small hawkmoths and up to 16 hours for large robust specimens. Alternatively, the vial containing the specimen in KOH can be heated in a water bath for a fast result, but care is needed to avoid macerating the genitalia too drastically. The genitalia should still have pigmentation and their scaling intact but be just soft enough to be manipulated, and all muscle tissue should be dissolved. (2) Wash the genitalia thoroughly to remove all traces of KOH (rinsing in dilute acetic acid can help) and transfer to a petri dish with water. (3) Remove abdominal sclerites and brush off the scaling from the valvae making sure to leave any stridulatory scales if present. If the scaling falls off in lumps then the preparation has been a little over-treated with KOH, and if the scaling cannot be removed the genitalia may need to be returned to KOH. (4) While still in water spread the valvae wide apart to expose the phallus and other structures previously hidden. On releasing pressure, the valvae may partly return to their closed position but if they cannot be forced apart, they will require further time in KOH. (5) At this point the phallus can be removed if desired. It is attached by a membrane near midlength. Pull the phallus towards the rear to expose the membrane that encircles the phallus in a tubular fashion. Tear this membrane with a probe, which will then allow the phallus to be withdrawn. This may require several attempts. Pulling backwards is usually best but 29/08/19 11:11:06.73 COLLECTION AND PRESERVATION sometimes armature at the apex of the phallus precludes this, and it then has to be removed by pulling forwards. (6) If the vesica is to be extruded, it must be done at this point before the dissection is transferred to alcohol or glycerol, which causes hardening. Using a very fine syringe inject water into the phallus base, inserting the point of the needle into the ductus seminalis. The pressure of the injected water should cause the vesica to evert from the phallus tip but this is not always successful. (7) For illustrative purposes, it is usual practice to present the genitalia with the valvae widely spread and the uncus and gnathos turned laterally. To turn the uncus and gnathos laterally snip one arm of the vinculum that supports them (usually the left side so that the uncus and gnathos turn to the right). The removed phallus must be illustrated separately at the same magnification. (8) The spread valvae and sidewise uncus and gnathos will not stay in their desired positions without fixing in absolute alcohol. To do this, position the genitalia as desired under a glass plate (excavated glass block covers are ideal) while still submerged under water. Prop the glass plate up a little on one side (a small nail is ideal for this) just sufficiently to slip the spread specimen underneath so that the glass plate prevents the valvae closing, then remove the nail to drop the glass plate. Sometimes extra weights may be needed to flatten the preparation. With the genitalia in position siphon off the water and replace it with absolute ethanol and leave to soak for an hour or two after which the genitalia will be hardened in the desired position. Male genitalia may be stored in 80% ethanol or mounted on a microscope slide. Female genitalia are treated similarly but because many diagnostic features are unsclerotised membranes, the process differs as follows: (1) Follow steps 1–2 as for male genitalia but remove the whole abdomen as the corpus bursae may extend full length and allow less time in the KOH solution. (3) Wash thoroughly and remove excess abdominal tissue being careful not to dislodge the terminal rod-like apophyses. Clear excess degraded muscle tissue to leave the corpus bursae clearly exposed for examination. (4) To assist in later diagnoses, it is often helpful to fully inflate and harden the bursa copulatrix 190613 Hawkmoths of Australia 3pp.indd 21 21 by injecting through the ductus bursae with absolute ethanol. (5) Store in 80% ethanol or glycerol or a mixture of both. The role of photography Photography is a very successful method to record hawkmoth life histories and habitats. Many quality digital cameras, available at reasonable price, have built-in macro functions and are capable of producing quality images of larvae, pupae, adults and habitats. For more enhanced images of hawkmoth eggs and first instar larvae, cameras with macro lenses and sophisticated flash arrangements are required. Any small camera or mobile telephone can produce adequate images of larger larvae, and the internet offers the opportunity for comparison with existing images through websites or contact with specialists who can assist with identifications. As a word of warning though, misidentification of sphingid images on the web, particularly of larval images, is commonplace. The compilation of an authoritative digital image library is an invaluable tool in the study of hawkmoths and the following procedures may prove helpful. Photograph the larva in situ (just as you discovered it) and without any type of flash, then in decreasing distance until the larva all but fills the frame. Next take several shots from slightly different angles in full sunlight and using flash, if available. These last images will maximise larval detail and provide the best opportunity for identifying unknown larvae based solely on the images. Finally, images should be taken of the foodplant on which the larva was found, in broad profile, the terminal section of a branch, and the flowers, fruit, or seeds, if any of these happen to be present. Such images are best supported by a pressed dried specimen of the plant for future identification and cross-referenced to the larval image. If at all possible, the larva should then be collected and kept until it has been identified with certainty; larval identifications are not always possible from images and may require rearing through to adulthood. The possibility always exists that the larva is one of the species whose life history is unknown and rearing such an unidentified larva through to adulthood then becomes a priority. 29/08/19 11:11:06.77 Rearing hawkmoths Collecting immatures Eggs from wild-caught females Due to the relative ease of collecting, most historical information that we have on Australian hawkmoths is based on adults attracted to light. Although these data form the basis of our taxonomic knowledge, they can provide little insight into the biology of the various species. Searching for eggs and larvae can often be very rewarding, especially for those species seldom encountered at light, such as species of Macroglossum, Cephonodes, Cizara and Angonyx. For example, Macroglossum prometheus lineata is rarely collected as an adult and is poorly represented in collections, but eggs and larvae are easily found on its larval foodplant, Morinda citrifolia, in coastal districts around Cairns and Mossman. Likewise, adult Cephonodes janus are seldom seen but eggs and larvae can be numerous on Psydrax odorata following the first heavy summer rains at The Caves north of Rockhampton. Similarly, the eggs and larvae of other so-called rare species can be found by searching the larval foodplants at the correct time of year. The characterisation of a species as rare should often be interpreted as a lack of knowledge of its biology and habitat preference, a subject discussed in detail by Kitching and Cadiou (2000). Although some species are infrequently observed and rare in collections, we consider no Australian sphingid species truly rare in nature. Searching for eggs and larvae offers opportunities for new discoveries and has at least five appealing advantages for the amateur or general nature lover: 1) no specialised equipment is required; 2) searching can be done during the day; 3) some species do not readily respond to light and rearing larvae is the best way to obtain specimens; 4) all aspects of the habitat can be enjoyed, most particularly the plants; 5) undamaged specimens are obtained for collections. Success in finding immatures requires knowledge of the local flora, either self-taught or through timely access to expert botanists. If the larval foodplant of a species is unknown in Australia, clues as to which plant family or genus is likely to be used here can be drawn from known foodplants of that species or a closely allied species in other parts of the world. Associating a larva with its foodplant offers the greatest opportunity for valuable new contributions, either by recording a new larval foodplant or, more significantly, discovering a previously unknown life history. While most larval searches are done during the day, some species effectively hide during daylight hours and only feed at night. These species often feed openly soon after dusk, and suspected foodplants can be located during the day and then checked in the early evening with a torch. Searching for the early stages of hawkmoths thus has the added benefit of leading to an enhanced understanding of their biology, as well as increasing the collector’s botanical knowledge. A keen enthusiast soon learns that a close relationship exists between some plant families and certain taxa of hawkmoths, with an individual species of hawkmoth often confined to a single plant species, genus or family (note Appendix 2). A recorded list of foodplants, together with other biological notes such as type of habitat, time of collection and instars present, is an invaluable tool in adding to our knowledge of hawkmoths. The inclination of females to lay eggs in captivity varies greatly among the sphingid genera. Females of Hopliocnema, Chelacnema and Coequosa readily lay eggs without the need to provide foodplant foliage or adult nectar sources. However, the females of most species are very reluctant to oviposit in captivity even if they carry mature eggs and are provided with larval foodplants and adult nectar sources within a flight cage. Best results with difficult nocturnal species are obtained in the field on the night of capture. Prior to beginning light trapping activities, in close proximity to the sheet, set up an enclosure of netting or very lightweight plastic, such as an oven bag (see below), over a living branch of the foodplant of the intended hawkmoth species. While males can be often far more common than females at light, when a targeted female does arrive, immediately place her in the enclosure. Such females will mostly already have fed and mated and are more inclined to lay. Once females are taken back to the lab, to increase chances of oviposition, they can be fed sugar water (10–20% sugar to water) or honey diluted with water, best prepared using sterilised containers and cooled boiled water. It is usually necessary to make females feed by uncoiling their proboscis so that its tip is immersed in the food solution, upon which the moth will usually start feeding. Yet, in spite of best efforts, results vary from individual to individual. As an example, over three years, we caged several dozen Psilogramma maxmouldsi with no results, only to have two females lay several dozen eggs each the following year, in spite of being treated in the same manner. 190613 Hawkmoths of Australia 3pp.indd 22 Rearing larvae Once eggs or larvae have been collected, successful rearing to pupation is dependent upon the ability to provide adequate supplies of healthy (not degraded or desiccated) foodplant. If the egg or larva was collected locally, this is not usually a problem (for foodplants from remote areas see below). Tubular netting sleeves can be placed over larvae on living branches, with the ends tied off. This approach is simple and effective across southern Australia as it provides good ventilation and light, some protection from parasitic flies and wasps, and allows for easy study of the developing larvae. However, in tropical and much of subtropical Australia, regardless of what steps are taken, almost nothing is safe from the persistent and voracious green tree ants (Oecophylla smaragdina). As a result, unless you are certain that green tree ants do not occur in the immediate area, we do not recommend sleeving outdoors in northern Australia. Two additional options to sleeving outdoors are commonly used. As an alternative, potted plants can often be purchased from native plant nurseries and sleeves placed over them in situations that are free of potential predators. To avoid larvae being exposed to systemic insecticides, pot plants should be purchased several months prior to being used and exposed to natural conditions to reduce the possibility of residual poisoning. Less expensive, but requiring more advance planning and preparation, plants can be transplanted from the wild or propagated for future use. The second alternative is to rear larvae indoors on foodplant cuttings. There are several variants of this approach. 29/08/19 11:11:06.82 REARING HAWKMOTHS Larvae can be reared in plastic food containers with a tight fitting lid, and we suggest this approach with all early instar larvae. The advantages of using an airtight plastic container, especially with early instar larvae, are keeping the foodplant from desiccating too fast, reducing the need for large cuttings, and allowing the use of small sprigs of the newest foliage which makes for safe, easy handling when checking on minute larvae. However, care needs to be taken to remove the larval frass and any excess moisture daily. Failure to do so can quickly lead to bacterial or viral infections, which are fatal in all cases. A base layer of absorbent paper toweling can be helpful in absorbing excess moisture and keeping the container clean if replaced daily. In some situations, paper toweling placed under the lid also helps by catching condensation forming there. Larvae can also be reared indoors by placing the base of the foodplant in water using a narrow-necked bottle and ‘sealing’ the neck with wads of cotton wool or paper to prevent larvae wandering down into the water. The larvae can then be contained by placing the bottle containing the foodplant in a cage or container. Polystyrene vegetable boxes are ideal for this purpose, as part of the lid can be cut out and replaced by gauze to offer ventilation and an observation window. Another variant is to use an oven bag (used for roasting) as a cover and tie it just above the point where the foodplant stem enters the neck of the water bottle. Oven bags are extremely lightweight, keep the foodplant in excellent shape for up to three times longer than most of the other methods, and have micro-perforations that all but eliminate moisture build-up. In addition, daily maintenance is very simple: the tie-off is removed, the foodplant is taken out of the oven bag, the bag is then inverted, the foodplant is once again enclosed and tied off. Cleanliness is of the utmost importance in rearing larvae. Ensure all breeding containers are sterilised before use. Avoid directly handling larvae, especially early instars, as they easily succumb to infection and/or poisoning, which can be passed on in this way. For example, handling larvae after having patted a pet with a flea collar or after having applied insect repellant while searching for larvae is inevitably fatal to the larvae. If larvae are to be handled always wash hands thoroughly beforehand. Artificial diets for larvae Many artificial diets have been developed for rearing lepidopteran larvae (e.g. McKinley 1971; Hervet et al. 2016) but few pertain specifically to hawkmoths. Gade (1980) developed one for Acherontia atropos, Kiguchi and Shimoda (1994) for Agrius convolvuli, Harbich (1994) for Hyles and Acherontia atropos, and Koch and Heinig (1977) for Daphnis nerii. Zagorinskii et al. (2013) provide a modified recipe for Acherontia atropos, A. lachesis and Daphnis nerii, and another for Agrius convolvuli and Hyles livornica. Retnakaran et al. (1985) provide a recipe for an artificial diet for rearing Daphnis nerii. Among the artificial diets developed for Manduca sexta over the years one of the more successful is that detailed by Ojeda-Avila et al. (2003). While that of Harbich (1994, reproduced in English by Hundsdoerfer, Tshibangu et al. 2005), is the simplest recipe and possibly suitable for many species, all others are complex and beyond the resources of most, so we refer only to the original publications. Some artificial diets are available commercially. Alternatively, Betts (2015) found that larvae of Hippotion celerio, Hyles livornica, Theretra oldenlandiae and Theretra alecto all thrive on hydroponically grown Iceberg lettuce, emphasising that it is essential that the lettuce has not been in contact with pesticides. 190613 Hawkmoths of Australia 3pp.indd 23 23 Housing pupae Just prior to leaving their foodplant to pupate many larvae change their appearance by turning a purplish brown or wine red. This is a telltale sign that the pupation process is about to begin. Reflecting its natural behaviour, the mature larva may then wander for several days in search of a suitable pupation site. Natural pupation sites of hawkmoths are varied. Many species pupate on the surface of the soil and are found within a cocoon composed of leaves, soil particles and other debris loosely bound by silk. In captivity, larvae of these species pupate on the base of their containers in the fashion described above if provided with a base layer of absorbent paper towel and dried leaves. The larvae of other species burrow into the soil to varying depths and make subterranean chambers in which to pupate. For these species, a container with at least 150 mm depth of potting mix or other friable soil is suitable. It is preferable that pupae are left in their underground chamber. The container should be supplied with vertical twigs or a gauze wall for the emerging moth to climb up on to expand its wings. Adults emerge from their pupa at species-specific times. Some hawkmoths tend to emerge shortly after dusk, others only much later at night, and others only in the early morning. Pupal duration is also highly variable but appears more dependent upon local seasonal conditions. During hot and humid periods, most hawkmoths will emerge two to four weeks after pupating. However, with the exceptions of Coequosa triangularis and C. australasiae, which overwinter as larvae, all other Australian sphingids pass through extended unfavorable weather conditions in the pupal stage. During very dry periods, pupae should be lightly misted occasionally with water to prevent desiccation. Rearing successive generations The challenge in rearing successive generations is obtaining matings. Not all species can be treated similarly and some degree of trial and error is usually needed. While species that do not feed as adults are easier to rear than those that need nectar, it appears that it is far easier to get bred adults to mate and lay in captivity than to use wild caught adults. Although there are no documented Australian examples, the following references report details of successful breedings that can be adopted either for a species also found in Australia or for related species. Betts (2015) successfully bred Hippotion celerio through multiple generations, and while he does not provide much detail, there seems to be sufficient information for others attempting to breed this apparently not particularly difficult species. Zagorinskii et al. (2013) reared multiple generations of Acherontia atropos (13 generations), Daphnis nerii (4 generations) and Hyles livornica (2 generations), and they give details for each species, including how they obtained matings and oviposition, and how they reared the larvae on artificial diets. Okelana and Odebiyi (2007) successfully bred Cephonodes hylas and provide considerable detail of how they caged and fed the adults and reared the larvae and pupae. Pittaway (1993) provides an excellent overview of hawkmoth breeding (pages 168–175) and gives notes for specific Western Palaearctic species (e.g. Agrius convolvuli p. 81, Acherontia atropos p. 83; Daphnis nerii pp. 116–117; Hyles euphorbiae, p. 137; Macroglossum stellatarum p. 134). He also summarises details on the many natural and artificially produced hybrids of hawkmoths (pages 64–66). Newman (1965) provides a report of his experiences in breeding some British species (pages 111–127), including how to feed adults and achieve pairing. 29/08/19 11:11:06.89 Biology Hawkmoths are essentially tropical insects and consequently most Australian species are found in the north, primarily in the wet tropics of Queensland. Sixty-four of Australia’s 87 species breed within 150 km of Cairns in northern Queensland. However, a few endemic species that breed only following opportunistic rains are found in arid regions of Australia. Many species are multi-brooded, especially in the tropics and in warmer and wetter areas of northern Queensland and the Northern Territory, where they can breed year round. Egg Most sphingid eggs are glossy, subspherical or nearly so, usually unmarked, and are usually light green or pale yellow. Notable exceptions are the grey eggs of Hopliocnema and Chelacnema, and the development with maturity of a bold, irregular reddish band along the circumference in some species including Psilogramma, Hopliocnema, Chelacnema and Coequosa. Eggs are usually laid singly on the underside of leaves of the foodplant, although sometimes on stems, tendrils or flower heads. Coenotes species are unique amongst Australian sphingids in that they lay their eggs in tight clusters. Females lay over several consecutive nights, sometimes visiting the same plant each night if oviposition sites are limited. Because some sphingid species share the same foodplant species, sometimes eggs of different sphingid species can be found on the same plant, particularly so on Vitaceae. Eggs of Australian species range in diameter from 1.0 mm for Macroglossum alcedo to 3.2 mm for Coequosa triangularis. They are usually proportional to the size of the adult but those of Agrius and some allied genera are unusually small. They are not constant in size, and the first eggs laid by a female tend to be larger than those laid towards the end of her life. The eggs of most Australian species hatch in 3–7 days although under excessive temperatures some can do so in 2 days, whereas incubation for eggs of Coequosa triangularis can take up to three weeks. None are known to overwinter. The total number of eggs laid by most species is 100 or more but the Australian endemic Hyles livornicoides has been found to lay as many as 460 eggs (N. McFarland pers. comm.). At the extreme, Okelana and Odebiyi (2007) recorded over 700 eggs laid by Cephonodes hylas under laboratory conditions. Those species that do not feed as adults tend to lay fewer eggs than those that do. Non-feeding species lack a developed proboscis and emerge with fully developed eggs, whereas those species that can feed have undeveloped gonads on emergence and must feed before the eggs can mature. Maturation of the eggs in feeding adults can often take several days. Larva Growth and moulting The larvae of most species pass through five instars. Amongst the Australian sphingids, Chelacnema ochra and Hopliocnema lacunosa are exceptions with four instars. Some large species, including Cerberonoton severina, have six instars and, unique among sphingids, the Australian endemics Coequosa australasiae and C. triangularis have seven or eight instars. Under unfavourable conditions such as temperature stress or inadequate food supply, some species can extend larval 190613 Hawkmoths of Australia 3pp.indd 24 development for one or sometimes more instars (Nijhout 1975; Jones et al. 1980; Davidowitz et al. 2003; Kingsolver 2007). Within the Australian fauna, additional instars have been observed in Agrius convolvuli, A. godarti, Cephonodes kingii and Coequosa australasiae. Kingsolver (2007) found that in the North American sphingid Manduca sexta, which normally has five instars, if adverse temperature and/or poor nutrition slowed growth in the early instars, those larvae had a high propensity for undergoing a sixth instar. Also he found that >95% of those larvae with a body mass >600 mg had only the typical five instars, whereas those with a body mass of <600 mg underwent a sixth instar. As speculated by Davidowitz et al. (2003), this suggests that, in M. sexta at least, a critical weight must be attained before pupation is triggered. Such variation in the number of instars is not confined to the Sphingidae, being widespread throughout insect orders (Esperk et al. 2007). Larval development (from eclosion from egg to prepupa) can be extremely rapid in some species, as short as 12 days for some Macroglossum species, for example, but most species usually take 3–4 weeks. Species of Coequosa are exceptions among sphingids in taking several months to reach maturity and it is the only sphingid genus in which the larvae overwinter, rather than the pupae. There is a tremendous increase in weight of the larvae from eclosion to maturity. For instance, Newman (1965) gives the weight increase in Sphinx ligustri as almost ten thousand fold. However, a considerable loss of weight occurs prior to pupation with the pupa usually weighing only about half that of the fully fed larva. Behaviour On hatching, the larvae of most species consume their egg shell, after which they rest for some hours before beginning to eat again. Usually they start with the leaf on which their egg was laid, either feeding from the leaf margin, especially on young shoots, or feeding on the leaf surface, thereby making holes in the leaf. Later instars always feed at the leaf margin. Many large larvae, especially when feeding on trees, work their way down the stem denuding it of leaves except for the projecting midribs, a behaviour known as ‘stemming’. Others consume the entire leaf, especially those feeding on understory and ground cover plants. Most larvae eat their larval skin at ecdysis but avoid the cast caudal horn. The larvae of most species feed at irregular intervals both during the day and at night. Some feed mostly at night, particularly the dark forms of Agrius convolvuli and Hippotion scrofa that are known to descend the foodplant to ground level where they rest during the day. Early instar larvae seldom venture far from their feeding site, usually resting along a leaf vein on the lower surface. Later instars hide amongst foliage, especially if green, although dark larvae of many species also hide in this manner. In addition to mottled colour patterns that help break up predator search images, the ventral surface of most last instar larvae is the darkest part of their body and the dorsal portion is the lightest. Larvae typically hang upsidedown, sometimes in an arched position, thereby directing the dark portion skyward which flattens their image, minimizes contrast and makes them even more difficult to detect among the plant shadows. 29/08/19 11:11:06.95 BIOLOGY When disturbed, the larger larvae of some species, especially in the subfamily Sphinginae, strike a posture reminiscent of the sphinx of ancient Egypt. This posture was first associated with the larvae of Sphinx ligustri by René Réaumur (1736) and was acknowledged by Linnaeus (1758), who described the genus Sphinx for the known hawkmoths of the time. Thus, the hawkmoths also became popularly called sphinx moths, although the purpose of the posturing is not clear. In a much more obviously defensive response, other larvae, when disturbed, regurgitate partially digested food that may contain toxins, as in the European Hyles euphorbiae, which feeds on toxic Euphorbia species (Hundsdoerfer, Tshibangu et al. 2005), and possibly the Australian hawkmoths Macroglossum vacillans and M. micacea micacea that feed on toxic Strychnos (Loganiaceae). The larvae of two desert dwelling species, Hyles livornicoides and Coenotes eremophilae, are renowned for their seemingly aimless wanderings across the ground, often in huge numbers, and they have become the subject of legends of local aboriginal tribes. Colour morphs and camouflage Almost all hawkmoth species have larvae in multiple colour morphs. The triggers for expressing these morphs are not well understood and very complex. Laboratory results from studies by Sasakawa (1973) and Sasakawa and Yamazaki (1967) with Cephonodes hylas and by Owen (1980) with Deilephila elpenor indicate that high population densities influence the expression of colour morphs. On the other hand, Grayson and Edmunds (1989) found that the intensity of the light reflected from the leaf and its surrounds determined colour in Laothoe populi and Smerinthus ocellata, particularly for the early instars. They also concluded that colour morphs in these species are, in part, inherited. Several additional factors have been shown to influence the expression of larval colour morphs in the sphingids Amphion floridensis, Eumorpha fasciatus, Xylophanes tersa and Enyo lugubris (Fink 1989, 1995). Amphion floridensis, the primary subject of study, has green and pink colour morphs and the expression of the pink morph, which varied from 2% to 86% in 35 broods, was influenced by temperature, foodplant choice, and leaf colour but was not influenced by rearing density or photoperiod. Although the mechanisms that drive larval polymorphism are varied and not easily attributable, they clearly affect survival fitness of larval populations, especially with respect to pressures from predation. As an example, Gerould (1921) reported that in wild populations of Colias philodice (Pieridae) larvae feeding on the same foodplant, blue-green larval forms were almost always found by sparrows (Passer domesticus), whereas the green forms went undetected. However, effective crypsis is not just a simple matter of colour variation. Behavioral responses in conjunction with polymorphism provide a further element of protection from predators. Green larvae of A. floridensis, E. fasciatus and X. tersa (Fink 1989), Acherontia atropos (Sevastopulo 1971), Agrius convolvuli (Edmunds 1975) and Sphecodina abbottii (Heinrich 1979) tend to position themselves along the underside of a leaf of the foodplant, whereas alternatively coloured larvae tend to use stems or branches toward the foodplant’s interior. Polymorphism is a topic ripe for additional research. Foodplants Larval foodplants are many and varied. There are 150 genera in 48 families used by the 87 Australian sphingid species. Some species are known to feed on just one plant species whereas others feed on a wide range of families, with the 190613 Hawkmoths of Australia 3pp.indd 25 25 Australian endemic Psilogramma casuarinae, for example, recorded from 35 plant species in 6 families, 26 of which are exotic. In contrast, another Australian endemic, Coenotes eremophilae, feeds on 30 plant species in 14 families, but only 5 are exotic. Rubiaceae and Vitaceae are the plant families most widely used as foodplants, collectively by 35 of the 87 Australian sphingid species. The most common foodplant is Cayratia clematidea (Vitaceae), used by 11 hawkmoth species. All currently known sphingid-foodplant associations in Australia are included in Appendix 2. Some foodplants, such as the Strychnos species used by Macroglossum vacillans and M. micacea micacea, are extremely toxic to vertebrates. Surprisingly, the potentially deadly barbed stinging hairs of the stinging trees (Dendrocnide species) seem not to harm the larvae of Theretra queenslandi. They seem not only to avoid being stung but devour the leaves with vigour. The larvae of Macroglossum dohertyi doddi are unusual in that they feed on the leaves of ant plants that have a symbiotic relationship with ants living within their bulbous bases. These carnivorous ants aggressively attack intruders but normally ignore the M. dohertyi doddi larvae that appear to be of no benefit to the ants. Pupa Nearly all sphingid species overwinter as pupae. The only known exceptions are the Australian endemic Coequosa species that overwinter as larvae. During very hot weather pupal duration can be as short as five days, as recorded for Macroglossum corythus approximans, and gradually lengthens into a few weeks until winter diapause is reached. Some species inhabiting arid regions, including Zacria vojtechi, Chelacnema ochra, Hopliocnema species and Hyles livornicoides, appear to diapause as pupae during dry years and await drenching rains before emerging, often in large numbers. Adults of nocturnal species emerge at species-specific times of the night, some shortly after dark, others much later at night, and others only in the early morning, therefore delaying their maiden flight until the second night. Diurnal species such as Cephonodes often emerge soon after dawn and fly the same day. Many species, particularly those in the subfamily Macroglossinae, pupate at ground level below a covering of debris where they spin an open net-like cocoon intertwined with dead leaves, twigs and grit. Others pupate in a subterranean chamber with its walls partly hardened by regurgitations from the larva. These chambers are often several centimetres below the surface and sometimes as deep as 20 cm or more. Most desert-dwelling species pupate subterraneously, including the Australian endemics Zacria vojtechi, Hyles livornicoides, Chelacnema ochra, Synoecha marmorata and species of Hopliocnema. Subterranean pupation may provide protection against desiccation in an arid environment in which species may need to remain as pupae for more than one season due to sporadic rainfall. In captivity, all species of Psilogramma and Coequosa pupate in deep subterranean chambers and use two adult emergence strategies. Some individuals work their way upward as a pupa and the adults only emerge from the pupal shell once at the surface, whereas others emerge from the pupa underground and the adults make their way upward to the soil surface while their unexpanded wings are still soft and pliable. Soil types and moisture levels in the ground may determine how emergence occurs in these and other species using subterranean pupation, but this process requires a great deal more research. Hippotion velox and some other species not found in Australia pupate arboreally amongst leaves of their foodplant. 29/08/19 11:11:07.03 26 HAWKMOTHS OF AUSTRALIA It is unclear why H. velox should pupate arboreally though it sometimes pupates in the leaf litter. Macroglossum dohertyi doddi normally pupates in concealed locations on the tree supporting its epiphytic foodplant, not entirely unexpected as these trees grow in swampy areas that are often inundated. In Australia, Theretra silhetensis intersecta feeds on Ludwigia species and Hippotion johanna feeds on Persicaria decipiens, plants that are semi-aquatic and at times inundated by water. The fate of most inundated larvae is unclear, but larvae of T. silhetensis intersecta and H. johanna have been observed wriggling across short stretches of slow moving water and then climbing adjacent plants on higher ground (JPT unpubl. obs.). Similar observations have been made of Deilephila elpenor (Linnaeus) in the western Palaearctic (Albin 1720; Newman 1965; Pittaway 1993). Just prior to pupation, green larvae of most species become predominantly pale wine-red and the pulsation of the heartbeat becomes highly visible along the middorsal line (Pl. 56, fig. h). This change in pigmentation first appears dorsally and spreads to a lesser degree laterally and sometimes ventrally within 24 hours. When ready to pupate larvae of some species wander considerable distances in search of a suitable pupation site; in fact, it seems as though wandering is a necessary prerequisite for pupation in some species of Agrius, Psilogramma and others. Characteristic larval markings remain visible on the pupal skin immediately after shedding the larval skin but they quickly vanish as the teneral pupa darkens to become brown. In those species with a very long proboscis, this is partly accommodated within a free sheath extending from the pupa as in Agrius, Cerberonoton and Psilogramma. Upon shedding of the larval skin in these genera, the proboscis is no more than a small rounded protrusion on the head of the forming pupa but it gradually grows longer, coiling if necessary, over an hour or so until fully developed before the pupa hardens (Pl. 14, figs g–l). Adult Feeding and pollination Most species feed by hovering in front of flowers as they take in nectar through their proboscis, usually at dusk or dawn. This behaviour has led to the popular name hawkmoths, as, while feeding, the moths can hover in a stationary position like a hawk. Macroglossum species have become popularly known as hummingbird hawkmoths because of their resemblance to feeding hummingbirds. Due to their yellow and black banded abdomen, transparent wings, and daytime nectaring, Cephonodes species are commonly called bee hawkmoths. Hawkmoths are specialist feeders of long tubular flowers although they may also feed on others, and they are some of the few insects capable of feeding while hovering in a vertical position. The classic example is the Madagascan orchid Angraecum sesquipedale with a tubular flower some 25 cm long pollinated by Xanthopan morganii praedicta Rothschild and Jordan, with one of the longest proboscises of any insect. Adult sphingids often develop feeding patterns where they visit the same flowers at the same time each day (Pittaway 1993). Australian species can be often found feeding at the flowers of Lantana, spider lilies (Hymenocallis littoralis), Ixora, Pentas, blue snakeweed (Stachytarpheta cayennensis) and male pawpaw (Carica papaya). Williams and Adam (2010: 26) mention that the lily Crinum pedunculatum is principally pollinated by hawkmoths (especially Theretra nessus, G. Williams pers. comm.). Hopper (1980) recorded five species of hawkmoths feeding at the flowers of the tropical rainforest tree Syzygium tierneyanum 190613 Hawkmoths of Australia 3pp.indd 26 (Myrtaceae) in northern Queensland. In the far north of Western Australia and the Northern Territory, hawkmoths are pollinators of boab trees, Adansonia gibbosa (Malvaceae) (Baum et al. 1998; Groffen et al. 2016). Armstrong (1979) lists five species of hawkmoths (previously recorded by A.G. Hamilton) visiting Clerodenrum tomentosum. Agrius convolvuli is one of the few pollinators of jacaranda in southern Australia (unpubl. obs. DAL, MSM). However, a few Australian genera, such as Chelacnema and Hopliocnema, have species with undeveloped mouth parts and are unable to feed. Species that feed are long-lived, often for six weeks or more, whereas those that cannot are short-lived surviving just a week or so. A few species, such as the European Macroglossum stellatarum, overwinter as adults and can live for many months. Migration Hawkmoths are strong fliers and well known for their ability to migrate. Agrius convolvuli migrates annually from northern Africa and the Middle East into higher latitudes, often as far north as the Scandinavian countries. While migratory patterns in Australia are still little understood, evidence suggests that Agrius convolvuli, A. godarti, Hippotion velox, H. scrofa, Hyles livornicoides, Macroglossum joannisi and M. vacillans are all migratory at times, often in large numbers and sometimes reaching far beyond the mainland to offshore islands. However, there is no evidence of migration north out of Australia into Indonesia or New Guinea. These sphingid migrations are not regular occurrences and why they take place is not clear, as dispersal is often into regions where there are no larval foodplants. Migration may be in part responsible for species temporarily establishing in marginal regions of their range during favourable years. Sexual communication and mating Females produce pheromones that males detect with their antennae, often at great distance, although much of the information about their range is speculative. For some species, this primary attraction may lead directly to a successful pairing. However, it is not always so simple and for many species pheromone attraction only brings conspecific adults into proximity. In some species, the males also produce pheromones as an aphrodisiac to subdue the female prior to copulation. Males of other species court the female by sound produced by stridulatory scales on their genitalia. As a consequence of these secondary male responses, Kitching and Cadiou (2000) suggest females may be quite selective in their choice of mates. Each species has its own flight activity window that efficiently facilitates male-female interaction, with some species pairing soon after dusk, some later at night or after midnight and some at dawn or shortly thereafter. Successful pairings may last as little as 30 minutes or may continue for hours. While coupled, the sexes remain stationary, joined back to back (Pl. 21, fig. m), the female at the top clinging to a branch or other vegetation while the male hangs below. Sight and ultraviolet light As in other moths, ultraviolet light is included in the visual spectrum of hawkmoths. As a result, flowers may look very different to them than they appear to our eyes. They are thought to use the horizon (the brightest part of the night sky apart from the moon) for orientation. An encounter with a bright light that out-competes the horizon causes them to become disoriented and fly towards the light source (Zborowski and Edwards 2007). While most species are readily attracted to light, there are some reluctant visitors to the light 29/08/19 11:11:07.10 BIOLOGY sheet such as Cizara ardeniae, and some rarely encountered species, such as Macroglossum prometheus lineata, despite both being relatively common species. In the non-arid regions of Australia, some species may respond to light sporadically or in small numbers prior to midnight, but numbers often increase as the night progresses. In contrast, most desert species fly primarily before midnight, often soon after dusk. Resting and camouflage Most Australian hawkmoths rest with their wings held flat in a ‘V’ shape but some, including Agrius and Acherontia in the tribe Acherontiini, hold their wings more tent-like over the body. Some Smerinthini not found in Australia adopt a rather different posture where the hindwings are brought forward and protrude in front of the forewings. In many species, the wings at rest provide cryptic patterns that assist in camouflaging adults during the day. They often rest against tree trunks or other substrates that match their appearance (Pl. 56, fig. l) especially in secluded dark spots, or amongst foliage where they hang by their front legs and resemble dead leaves. Hawkmoths as pests Larvae as pests In Australia, the larvae of 21 hawkmoth species have been reported as feeding on cultivated plants (Moulds 1981, 1984). Seven of these, Acosmeryx cinnamomea, A. miskini, Gnathothlibus eras, Hippotion celerio, Hyles livornicoides, Theretra latreillii and T. oldenlandiae attack grape vines, occasionally causing defoliation. Sweet potato vines are eaten by four species, Agrius convolvuli, Hippotion celerio, H. scrofa and Gnathothlibus eras, which sometimes cause notable damage. Psilogramma casuarinae is occasionally found on olives, Hippotion celerio occasionally on rhubarb, and there are several records of Coequosa triangularis on macadamias. Cephonodes cunninghami is a minor pest of coffee in Australia, and the closely allied species C. hylas is regarded as a major pest of coffee in Africa and much of Asia (e.g. in Nigeria, where it is responsible for 50–70% yield loss (Okelana and Odebiyi 2007)). Eleven species of hawkmoths feed on some 70 species of garden ornamentals in Australia (Moulds 1984), but rarely is damage significant. Prominent among them are Agrius convolvuli favouring morning glory; Gnathothlibus eras frequently found on ornamental grapes and pentas; Cephonodes kingii, C. australis and C. cunninghami on gardenia; Psilogramma casuarinae on privet and jasmine; Theretra oldenlandiae often on balsams and fuchsias; and Theretra latreillii also on balsams. Although adults of Hippotion celerio and H. scrofa are often very common and occur throughout most of Australia, their larvae are not common on garden ornamentals despite having been recorded on many of them. 27 that the three Acherontia species may each be specialised for entering the hives of different Apis species (for a full discussion see ‘Genus Acherontia’). To date, A. lachesis has been reported robbing only Apis dorsata, and consequently its introduction to the Australian mainland would not impact the Australian honey industry, but there is insufficient evidence to definitively say it could not rob Apis mellifera were it to establish itself on the Australian mainland. However, although Acherontia lachesis adults are too large to steal honey from modern commercial beehives with narrow slit entrances some caution is required should disease or varroa mites establish in Australia. Then, if Acherontia lachesis was capable of robbing Apis mellifera, it would have the potential to spread disease via feral colonies or through failed feeding attempts at commercial hives. Hawkmoths as human food and medicine The larvae of Hyles livornicoides and Coenotes eremophilae were eaten by Australian aborigines in the deserts of central Australia (Yen 2005). These larvae, known respectively as Yeperenye and Anumara by the Arrente people near Alice Springs, are exceptionally numerous after rain. They were kept alive until their gut cleared and then cooked in coals (Tindale 1972). Further details on cooking lepidopterous larvae from tribes in the Lake Eyre Basin given by Chewings (1936) suggest that the heads were removed before cooking and the larvae then either eaten or stored dry. Later they were ground, then kneaded into a paste and baked. Hawkmoth larvae are eaten in other parts of the world, especially in Africa and Asia. Usually the larvae and pupae are consumed. Yen (2015) notes that Clanis bilineata is farmed in China for food. Umermura (1943) lists Psilogramma increta as being wholesome food in Japan. In Africa, Silow (1976) records hawkmoth larvae being eaten by the Mbunda people of Zambia. In Botswana, Moreki et al. (2016) record Agrius convolvuli and Lophostethus dumolinii as human food that is cooked by boiling, roasting or frying. These two species were by far the most consumed insects in the two villages studied. The larvae of L. dumolinii are consumed by first removing the head and clearing the gut prior to cooking and the bodies then dried in the sun for later consumption. It was not stated what stages of A. convolvuli were eaten. In the deserts of North America, some native tribes ate the larvae of Hyles lineata as a seasonal staple (Brown 1967). Hawkmoths have also been credited with therapeutic qualities. The larvae of six species, Acherontia styx, Agrius convolvuli, Macroglossum stellatarum, Deilephila elpenor, Theretra nessus and T. oldenlandiae have at various times been considered to be effective in treatments of tuberculosis, stomach upsets, mumps, tumours, fever and snake bite (Umermura 1943; Schimitschek 1968; Meyer-Rochow 2017). Natural enemies Adults as pests Death’s head hawkmoths are the only sphingid species whose adults are considered pests because they are known for taking honey from bee hives. But these moths not only have the capacity to consume honey but are potential carriers of disease. Of the three species of death’s head hawkmoth, only Acherontia lachesis is likely to establish on the Australian mainland having recently spread through New Guinea to Torres Strait. The reputation of death’s head hawkmoths as pests is primarily based on observations made of A. atropos robbing Apis mellifera in Europe and northern Africa. However, Koeniger et al. (2010) has shown that A. styx and A. lachesis are also avid consumers of honey. Current observations suggest 190613 Hawkmoths of Australia 3pp.indd 27 Natural enemies play an important role in regulating the populations of insects, including Lepidoptera. Without them, insects would overrun the world because of their enormous reproductive potential. However, there have been very few studies of natural enemies and most have concentrated on insects of economic significance. The general characteristics of four broad categories of natural enemies, predators, parasites, parasitoids and pathogens, are discussed below, and a summary of all known parasitoid records from Australian sphingids is provided (Appendix 1). Predatory and parasitic attacks take place quickly, are seldom observed, and even less frequently recorded. Even when predatory attacks are observed, the destruction of the 29/08/19 11:11:07.18 28 HAWKMOTHS OF AUSTRALIA prey and quick retreat of the predator greatly reduces the opportunity to accurately determine the identity of one or even both. Parasitic attacks are similarly short but, in contrast, while the host is still present, it often bears little evidence of the attack. Parasitoid attacks also involve short-term initial encounters with the host, but if the host is collected, the presence of the parasitoid may eventually be revealed. Although it has been difficult or impossible to identify host larvae in the past, most Australian sphingid larvae can now be identified in later instars and relying on various parasitoid experts (Appendix 1) and the parasitoid images in this book, one can now have reasonable confidence in recognising both participants in most Australian parasitoid-sphingid larval associations. Identifying parasitoidsphingid egg associations is often far more difficult. A few sphingid species have unique eggs, which makes host identification easy. Among the many sphingid species with ‘typical’ green ovoid eggs, narrow foodplant associations and context often allow identification of host eggs to species. However, there are limitations, especially when the larval foodplant preference involves members of the grape family (Vitaceae). We have found as many as five sphingid species with ‘typical’ greenish eggs using multiple vitaceous plants at a single location. As a result, identifying individual sphingid eggs to species is impossible in these situations. Pathogens Pathogens, including viruses, bacteria and pathogenic fungi, infect hawkmoths. Viruses and bacteria are particularly common natural enemies of immature stages and highly contagious in confined environments. Viruses Nuclear polyhedrosis viruses (NPV) are members of the Baculovirus family. Many species infect insects, predominantly Lepidoptera larvae (Federici 2009). These viruses tend to be species-specific and are categorised using an acronym prefix to identify their host species (Capinera 2008b). Infection occurs when a larva ingests or contacts substrates contaminated by body emissions of infected larvae. On ingestion, viral replication begins in the midgut. Occlusion bodies are solubilized in the gut lumen, releasing occlusion-derived virions that are thought to enter the target epithelial cells by fusing with the plasma membrane at the cell surface. Eventually, the host liquefies from having its cells lysed by the virus, and the larval cadaver, usually hanging by its prolegs, ruptures spreading infectious occlusions (polyhedral bodies) in the nearby environment. Nuclear polyhedrosis viruses are found throughout the natural environment but are usually thinly spread so that they normally have little impact on wild Lepidoptera populations. Only when a host becomes sufficiently numerous does the virus have a noticeable impact. Bacteria While most lepidopteran-associated bacteria are beneficial in assisting in digestion and other bodily functions, a few are fatal, although mortality from bacterial infection in hawkmoths is uncommon. The most often encountered fatal bacterium is Bacillus thuringiensis, which comes in numerous species-specific subspecies and isolates infecting lepidopteran larvae (Maagt et al. 2001). This species-specific attribute has made B. thuringiensis the preferred bacterium for the development of many bio-pesticides (Federici 2009). During sporulation, B. thuringiensis produces both endospores and crystalline inclusions (Capinera 2008a). The endospores, resistant to environmental stress, allow long-term survival. 190613 Hawkmoths of Australia 3pp.indd 28 The endospores lead to infection when larvae eat infected leaves. Infected larvae cease feeding because the inclusions cause paralysis of the mouth and gut, the larva eventually succumbing to a general paralysis leading to death. Many arthropods, including adult hawkmoths, carry symbiotic bacteria of the genus Wolbachia within their reproductive systems. Though Wolbachia has been widely studied in other Lepidoptera families (Ahmed et al. 2015; Ilinsky and Kosterin 2017) little is known about it in hawkmoths. Wolbachia infection can be either beneficial or detrimental to the host. Some Wolbachia have been used as biological control agents but no studies involving hawkmoths are available. Entomopathogenic fungi Entomopathogenic fungi, also known as entomogenous fungi, infect the larvae, pupae and adults of insects. Only species of Cordyceps and Beauveria (Cordycipitaceae) are known to infect hawkmoths. Shrestha et al. (2016) lists six Cordyceps species that infect the larvae or pupae of 13 hawkmoth species, but none is an Australian record. Koval (1974) records C. albocitrinus from the larva of a sphingid and Liu et al. (1984) record C. taishanensis also from a sphingid larva, from Russia and China respectively. Changes to the International Code of Nomenclature for algae, fungi, and plants (McNeill et al. 2012) abandoned the system of having different names for the different reproductive stages of the same fungus. This has resulted in nomenclatural changes for many entomopathogenic fungi since 2012, including Cordyceps (Kepler et al. 2017; Mongkolsamrit et al. 2018). Fan et al. (2019) provide a phylogeny for many Cordycipitaceae that includes Cordyceps and Beauveria. The only larval record for Australian hawkmoths is of a suspected Beauveria sp. infecting Hippotion celerio (Pl. 29, fig. m), and there are no pupal records. There are three records of hawkmoth adults being infected in Australia by a Cordyceps sp. that is peculiar to adult moths (Hope 2002, unpubl. obs. MSM) (Pl. 6, fig. g). These fungi produce conidia (spores) on long filamentous structures known as conidiophores. On germination, the conidia produce resting spores that permeate the environment to infect the next generation of hawkmoths. These fungi require damp environments to reproduce and all three of the known specimens were found in temperate rainforests of the mid North Coast of New South Wales. Predators Predation events are seldom observed and little evidence of an attack is subsequently found, hence the significance of predators on sphingid populations is impossible to assess. However, several morphological and behavioural responses in Lepidoptera suggest that predation has an influence on populations. The two primary groups of vertebrate predators are birds and bats (Pl. 32, fig. l). To avoid diurnal predators, the adults of most hawkmoth species tend to remain inactive during the day, seeking sheltered locations, and, as indicated above, rely upon crypsis to avoid detection. As a secondary defence upon discovery, the adults of some species have cryptic forewings that conceal colourful hindwings that resemble large eyes. The sudden flashing of such hindwings may produce a startle effect that provides the moth with an escape opportunity. Finally, some sphingid adults have sharp tibial spines that may provide some defence against predators. Many birds are active caterpillar hunters. They form ‘search images’, but what is unclear is whether the images are limited to the caterpillar itself or focus on a broader context, such as the nature and condition of the foodplant. The ‘search image’ that we employ to search for larvae in the field relies more upon 29/08/19 11:11:07.24 BIOLOGY characteristic larval feeding patterns and, in some circumstances, locating frass, rather than the larva itself. Almost certainly, birds employ even more acutely developed ‘search images’ that appear to have influenced the larval feeding patterns of some Lepidoptera. Some larvae reduce the likelihood of bird predation by feeding at night, or by moving frequently among the branches of larger foodplants or from plant to plant, thereby minimizing characteristic plant damage (Heinrich 1979; Heinrich and Collins 1983). Most hawkmoth larvae have, to a varying degree, some form of countershading and/or dorsolateral striping that enhances the ability of the larva to blend into its natural surroundings. Intraspecific larval variation within a given population, the presence/absence of irregular patterns or blotches which often match the colour of the foliage and/or woody portion of the plant (Pl. 55, figs g, h, j), help to break up the larval profile in varying ways. Schmidt (1990) offers an insightful discussion of the advantages of larval polyphenism in the sphingid, Eumorpha typhon, especially when considered in the context of orientation within the shadows and subtle colour textures of their foodplants. As a result, multiple morphs within a population reduce the potential for a refined avian search image. Some sphingid larvae are well known for prominent eyespots that may deter birds (Pl. 64, fig. j). However, Collins and Wagner (2014) caution against perceiving such markings from a human perspective as they may not be the same as from a bird’s view, especially as birds can see ultraviolet light. Bura et al. (2016) found that some sphingid larvae can produce sound when harassed, which they suggest is also used defensively, either as a warning of a pending chemical defence or to startle predators. Nocturnal hawkmoths risk being preyed upon by owls and bats. Owl predation on adult hawkmoths is sometimes observed at lights and probably also occurs in natural settings. Many bats are insectivorous, and it is apparent that moths and bats have been involved in a co-evolving ‘arms race’ for a long period of time (Fullard 1998). Bats are extremely efficient moth predators and locate prey by ultrasound and echolocation. Some sphingids have sensory organs on their labial palps that enable them to hear and take evasive action in response to bat-generated ultrasound frequencies (Roeder 1974; Göpfert and Wasserthal 1999a). Three sphingid species, including Theretra nessus Drury which occurs in northern Australia, have been shown to produce ultrasonic signals in response to bat echolocation (Barber and Kawahara 2013; Kawahara and Barber 2015). Males of some species produce the sounds by grating specialised scales on the genital valvae against the last abdominal segment, while the source of the sounds has yet to be identified in females (Barber and Kawahara 2013; Kawahara and Barber 2015). However, males of Macroglossinae may instead rub the valve scales against each other to produce the sound (I.J. Kitching pers comm.). Recent experimental results indicate that these sounds serve primarily as a jamming function in response to bat echolocation (Kawahara and Barber 2015). In addition to the impact of vertebrate predators, anecdotal observations suggest that in some settings invertebrate predation of sphingid eggs and larvae is far more significant. Sphingid larvae are subject to attacks by ants, particularly green tree ants (Oecophylla sp.) and bull ants (Myrmecia sp.) (Formicidae), as well as by spiders (Pl. 70, fig. i), assassin bugs (Reduvidae) and carabid beetles (Pl. 17, fig. f), and nymphs of erythraeid mites attack sphingid eggs. Parasites The term parasite is often incorrectly applied to the various flies and wasps that emerge from immature stages of other 190613 Hawkmoths of Australia 3pp.indd 29 29 insects. Such flies and wasps are properly termed parasitoids. While parasitoid adults are independently free-flying, their larval stages are totally dependent upon their host, both for food and shelter, and their long-term presence almost always directly leads to the death of the host insect. Conversely, except for microbes which are here treated separately, a true parasite derives temporary (see exceptions below) food and, in some cases, shelter at the expense of its host and this parasite-host interaction rarely leads directly to the death of the host. However, such attacks may lead to a weakening of the host making it more susceptible to predators, fungal, bacterial or viral infection, either indirectly through the opening of a wound or through direct transmission by the parasite as a vector. Compared to the large number of parasitoid-sphingid associations reported, there are few parasite-sphingid relationships documented in the literature. Globally, the reported relationships include nematodes (Nematoda: Acugutturidae), mites (Acari: Erythraeidae and Otopheidomenidae) and midges (Diptera: Ceratopogonidae). Brief discussions are included here to alert readers to their potential occurrence in Australia. Nematodes In the tropics of the Americas, three genera (Acugutturus, Vampyronema and Noctuidonema) in the nematode family Acugutturidae have been identified as external parasites on the adults of six families of Lepidoptera, including two sphingid genera (Rogers et al. 1990; Marti et al. 2000; Marti et al. 2002). These nematodes feed using a long, piercing stylet and while the interaction is debilitating for the host, it is not usually fatal (Simmons and Rogers 1996). There are no known associations of Sphingidae with nematodes in Australia. However, two of the large mermithid nematodes (Mermithidae), Amphimermis bogongae Welch and Hexamermis cavicola (Welch), attack bogong moths (Agrotis infusa Boisduval: Noctuidae) in their summer aestivation sites (Common, 1954; Welch, 1963), so there is the possibility that nematode-sphingid associations may be found in Australia. Mites The association between mites and Lepidoptera is an extremely old one that has been preserved in the fossil record. Mites in the family Erythraeidae have been found on gracillariid and tineid moths preserved in amber (Poinar et al. 1991). Erythraeid mites may act as either predators or parasites, although the distinction is difficult to discern in some cases. In Australia, erythraeid mites have been found associated with the adults and larvae of several lepidopteran families but there are no reported associations with sphingids as yet (Southcott 1966, 1972, 1991; Treat 1975). However, erythraeid mite nymphs in the genus Charletonia are predators on lepidopteran eggs, including Macroglossum errans in Tolga, Queensland (G. Sankowsky pers. comm.). Mites in the family Otopheidomenidae are commonly associated with adult Sphingidae in the Neotropics and the genus Prasadiseius exclusively so (Prasad 2011a, 2011c). Numbers on individual hawkmoths can be very high (Prasad 2011b). However, even when hundreds of mite eggs, nymphs, adults and fecal matter are found on a single host, this is not definitive proof of a parasitic relationship. Prasad (1970) indicated that even under microscopic examination, lesions of the host exoskeleton or other damage to the host could not be found. Halliday (1994) first reported otopheidomenid mites from Australia. The mites were associated with Hemiptera and, as with the Neotropical sphingid associations, all life cycle stages 29/08/19 11:11:07.31 30 HAWKMOTHS OF AUSTRALIA were present but parasitism could not be conclusively confirmed. However, there is a significant morphological character to consider when assessing the nature of the otopheidomenid mite-moth relationship. The chelicerae of otopheidomenid mites are long styli that would be effective for piercing and sucking, whereas predatory mite families have biting mouthparts (De Lillo et al. [2002]; Prasad 2012). This morphological adaptation and the presence of the entire life cycle on their host strongly suggest that otopheidomenid mites are feeding upon the haemolymph of their hosts, and hence are parasitic. It should be noted that Prasad (2011c) identified the Neotropical sphingid associations by examining preserved museum specimens. So far, very limited and random spot checks of museum specimens have failed to establish mitesphingid adult associations in Australia. Midges (Diptera: Ceratopogonidae) De Geer (1752) was the first to report midges sucking haemolymph from lepidopteran larvae. Although he did not specify which larvae were involved, he did characterise the host as ‘Nos grandes Chenilles’ [our large caterpillars]. Based upon the European Lepidoptera fauna, there is a possibility that his observations involved sphingids as hosts. Worldwide, the midge genus Forcipomyia has been reported as parasitic on the adults and/or larvae of at least eight families of Lepidoptera (Moore 1958; Debenham and Wirth 1984; Debenham 1987; Kawahara et al. 2006; Salvato and Salvato 2008; Koptur et al. 2013). Only females are known to feed, which suggests that host haemolymph is necessary for their reproductive cycle. There are no midge associations previously reported with adult sphingids in Australia, but Forcipomyia nr. proximornata (A. Borkent pers. comm.) are known to attack the larvae of three species: Agrius convolvuli (D. Davey pers. comm.), Psilogramma menephron nebulosa (B.M. Fjellstad pers. comm.) (Pl. 57, fig. j) and Theretra latreillii (B.M. Fjellstad pers. comm.). Moore (1958) reported midge feeding on limacodid larvae in NSW and indicated that the larvae died from a suspected viral infection soon after. Previously, Mayer (1955) had reported that a large caterpillar population in Cuba that was attacked by midges died of an apparent viral disease. Much later, Wirth (1972, 1975) speculated that the feeding habits of female midges were consistent with a vector capable of spreading polyhedral virus and other diseases. However, most hosts are unaffected; a Psilogramma menephron nebulosa larva attacked by three midges simultaneously pupated successfully and a healthy adult emerged (B.M. Fjellstad pers. comm.). Parasitoids (Appendix 1) Parasitoid-host interactions can be highly complex. Parasitoids use one of two strategies while feeding upon the haemolymph and fatty tissues of the host. Idiobiont parasitoids arrest the development of their host at the time of oviposition, thereby relying upon a fixed host biomass, as in the case of the various microwasps that attack sphingid eggs. In contrast, following the initial attack, koinobiont parasitoids allow their host to continue developing through subsequent larval instars and, in some cases, even through ‘successful’ pupation, until the growing parasitoid eventually kills it. This strategy provides a greater food resource for the parasitoid larva and is employed by all tachinid flies and some wasps. Koinobiont parasitoid strategies are then further subdivided. All tachinid flies and many wasps undergo their entire larval development within the body of the host and are called endoparasitoids, whereas some wasps spend their larval development attached and fully exposed on the outer cuticle of the host and are called ectoparasitoids. 190613 Hawkmoths of Australia 3pp.indd 30 Many wasps and all flies that attack Sphingidae are primary parasitoids; the sphingid larva provides all of the food and ‘shelter’ necessary for the parasitoid larva or maggot to complete development. However, not all parasitoid-host relationships are so straightforward, and many parasitoids are themselves subject to parasitoidism. When a primary parasitoid is attacked by a secondary parasitoid, the relationship is referred to as hyperparasitoidism. All hyperparasitoids are wasps. There are two types of hyperparasitoidism, obligatory hyperparasitoidism in which the secondary parasitoid always relies upon a primary parasitoid as its host, and facultative hyperparasitoidism in which the secondary parasitoid acts usually as a primary parasitoid (in sphingid larvae or eggs) but may opportunistically attack another primary parasitoid. To date there are no records of hyperparasitoidism associated with sphingids from Australia. However, given the number of such relationships documented from other parts of the world (Noyes 2017), closer scrutiny may well uncover them. Hymenopteran (wasps) and dipteran (flies) parasitoids play an important role in keeping insect populations in balance. Parasitoid wasps of various families attack the eggs, larvae and pupae of their sphingid hosts, whereas parasitoid flies attack the larvae and purportedly also pupae. As a natural control, hymenopterous egg parasitoids greatly reduce population numbers and larval parasitoids (wasps and flies) further reduce those numbers. Hymenoptera The suborder Apocrita in the Hymenoptera consists of wasps, bees, and ants. Eleven superfamilies within the Apocrita are Lepidoptera parasitoids and play a vital role in holding natural insect populations in check. Among them, a number of families contain biological control agents important for the agricultural economies of the world. The taxonomy of parasitoid families is poorly known and, except for economically important species, host associations are mostly unknown. As an example of the taxonomic work still to be done, it is estimated that there are somewhere between 60 000 to 500 000 species in the superfamily Chalcidoidea; however, only some 20 000 species have been named (Noyes 2017). Most Hymenoptera have a female sex ratio bias and fertilised eggs become females and unfertilised eggs become males (arrhenotokous parthenogenesis) (Heimpel and de Boer 2008). However, examples in which females are produced from unfertilised eggs (thelyotokous parthenogenesis) have also been documented. Both sex determination mechanisms occur among the apocritan parasitoid superfamilies Chalcidoidea, Ichneumonoidea and Platygastroidea (Cook and Crozier 1995). Parasitoids in these superfamilies can employ idiobiont or koinoboint strategies and be ectoparasitoids or endoparasitoids. It is generally agreed that endoparasitoid-host relationships require a great deal of specialisation and may have driven super-speciation among endoparasitoids (Sharkey 2007). Egg parasitoidism by chalcidoid and platygastroid microwasps has a significant impact on sphingid populations. High levels of egg parasitoidism have been reported in many lepidopterous families, and specific sphingid examples include a 93.8% parasitoidism rate on Daphnis nerii on Guam (Moore and Miller 2008) and a 98.5% rate on Xylophanes pluto in Florida (Tuttle 2007). We present here the first parasitoidsphingid egg associations from Australia. Among the 19 superfamilies of the Apocrita recognised worldwide, three superfamilies and eight families associated with Sphingidae have been reported from Australia to date (see Appendix 1), namely Chalcidoidea (Chalcididae, Encyrtidae, Eulophidae, Eupelmidae and Trichogrammatidae); 29/08/19 11:11:07.38 BIOLOGY Ichneumonoidea (Braconidae and Ichneumonidae) and Platygastroidea (Scelionidae). However, the paucity of records is presumably the result of insufficient reporting and additional families are almost certain to occur here. Each of these families is detailed below. Chalcididae (Chalcidoidea) The Chalcididae are a comparatively small group of wasp parasitoids with five subfamilies (Chalcidinae, Dirhininae, Epitraninae, Haltichellinae and Smicromorphinae) comprised of 87 genera and approximately 1464 species (Noyes 2017). There are 22 genera and 178 species in Australia (Australian Biological Resources Study 2009). There were no reported chalcid associations with sphingids (Noyes 2017), but Gnathothlibus eras and Macroglossum vacillans eggs from northern Queensland recently yielded undetermined Brachymeria. There are 59 Brachymeria species currently recognised from Australia (Australian Biological Resources Study 2009). Encyrtidae (Chalcidoidea) The Encyrtidae are important biological control agents, particularly for scale insects. Noyes (2017) indicated that there are 54 genera in Australia but did not report any associations with Sphingidae. However, he did note that Agrius convolvuli (Linnaeus), whose cosmopolitan range extends throughout much of Australia, is host to Ooencyrtus in other parts of the world. In addition, Cephonodes hylas (Linnaeus), which was previously treated as a member of the Australian fauna and is closely related to C. australis Kitching and Cadiou, is host to two Ooencyrtus species (Noyes 2017). Sixteen Ooencyrtus species are recorded from Australia (Australian Biological Resources Study 2009). Rearings from our current fieldwork include two and perhaps three Ooencyrtus species associated with the eggs of Australian sphingids (Appendix 1). As the generic name suggests, Ooencyrtus is known primarily as an egg parasitoid and multiple wasps usually emerge from a single host egg (Pl. 1, figs a, b). No examples of hyperparasitoidism by, or on, Ooencyrtus have been reported from Australia although examples of both exist in other parts of the world (Noyes 2017). Ooencyrtus species have been reared extensively as biological control agents. In spite of considerable laboratory rearing, it is unclear whether there is a sex bias. Studies by Wilson and Woolcock (1960) and Wilson (1962) were inconclusive. Three of our five Ooencyrtus-sphingid collections in nature did not include males, but our Macroglossum prometheus lineata Lucas sample comprised 5 females and 1 male and our Nephele subvaria (Walker) sample 9 females and 3 males. The N. subvaria sample, in particular, suggests that males may not be as scarce in nature as previously reported. Several observations on one of the most widely used species, O. kuvanae (Howard), are worth noting. Hofstetter and Raffa (1998) reported that O. kuvanae females seeking host eggs are attracted to residual volatiles associated with the ovipositor of the host female. In addition, the age of the host eggs upon discovery impacted not only the number of parasitoids but the gender ratio; older eggs resulted in fewer parasitoids and higher male to female ratios than younger eggs. And finally, they reported that the number of offspring per female and the proportion of female offspring are inversely related to the density of the local adult wasp population. Eulophidae (Chalcidoidea) Eulophidae number approximately 115 genera and 850 named species (Noyes 2017). Although Erebidae, Geometridae, Gracillariidae, Nolidae, Noctuidae, Notodontidae, Pyralidae, 190613 Hawkmoths of Australia 3pp.indd 31 31 and Tortricidae have been previously documented as hosts (Noyes 2017), associations with Australian Sphingidae have not been previously reported. The only known eulophid-sphingid associations are with the genus Euplectrus, which has 18 named species in the Australian fauna (Australian Biological Resources Study 2009). While there are no reports of an Australian Euplectrus acting as a hyperparasitoid, an undetermined Euplectrus and E. bicolor (Swederus) have themselves been parasitised by eulophid wasps, Pediobius bruchicida (Rondani) and P. atamiensis (Ashmead) respectively (Berry and Mansfield 2006; Noyes 2017). Euplectrus species are primary ectoparasitoids of lepidopterous larvae (Pl. 3, fig. c; Pl. 4, fig. i; Pl. 33, fig. e). Jones and Sands (1999) reported that eggs are primarily deposited on the cuticle of 2nd and 3rd instar larvae of fruit-piercing moths (Erebidae) and that parasitoid survival rates are significantly reduced when late instar host larvae are attacked. Yet 1st instar larvae are also attacked by a closely related Euplectromorpha species (Pl. 62, fig. b). Once Euplectrus larvae have completed development, they anchor the mummified host carcass to a leaf or to debris with a lattice of silk and then spin dense silken cocoons within that silk network (Pl. 4, fig. j, Pl. 33, fig. f). Eupelmidae (Chalcidoidea) The Eupelmidae is the least speciose of the microwasp families associated with sphingids in Australia. There are three subfamilies (Calosotinae, Eupelminae and Neanastatinae), 45 genera and 907 species worldwide (Noyes 2017). There are 13 genera and 175 species reported from Australia (Australian Biological Resources Study 2009). Anastatus is the only genus of the family reported in association with sphingids in Australia. Its species are among the largest of the egg parasitoids and attack a number of insect orders, but in spite of the large number of species recognised worldwide, relatively little is known about them. The adults are sexually dimorphic to such a degree that in relatively few cases have males and females been conclusively associated (Pl. 1, figs c–e). Males are seldom reported and many species are known only from females. Not surprisingly, given their relatively large size in relation to the size of their host eggs, all known Anastatus-Lepidoptera associations involve a single parasitoid per host egg. However, Anastatus can also act as facultative hyperparasitoid; three species (A. bifasciatus, A. pearsalli, and an undetermined Anastatus) have been documented as hyperparasitoids of braconids (Muesebeck and Dohanian 1927: 24–25; Berry and Mansfield 2006) and dipterans (Clausen 1940: 192). In addition, Anastatus can itself be the victim of hyperparasitoidism with one Australian example, a hemipteran as the primary host and A. biproruli Girault as the secondary host being attacked by Centrodora darwini Girault (Aphelinidae) (Noyes 2017). Thirty-seven Anastatus species are known from Australia, but only a few have been associated with hosts (Australian Biological Resources Study 2009; Noyes 2017). Although sphingid eggs serve as primary Anastatus hosts in other regions, the only previously known lepidopteran association in Australia was with a species of Notodontidae (Noyes 2017). Four Anastatus-sphingid associations are now known from Australia (Appendix 1). Trichogrammatidae (Chalcidoidea) Trichogrammatids are primary parasitoids of insect eggs although occasionally they act as facultative hyperparasitoids (Strand and Vinson 1984). There are approximately 80 genera and 850 named trichogrammatids worldwide plus an estimated 4000 unnamed taxa, with 32 genera and 170 species named from Australia (Noyes 2017). Worldwide, 29/08/19 11:11:07.45 32 HAWKMOTHS OF AUSTRALIA Trichogramma is the genus most frequently associated with Sphingidae. Except for the introduction of T. pretiosum Riley and T. carverae Oatman and Pinto to control an outbreak of Hippotion velox (Fabricius) on several Great Barrier Reef islands (Smith et al. 2004), there were no previously recorded trichogrammatid-sphingid associations known from Australia. However, 10 additional associations are now reported (Appendix 1). The taxonomy of the family Trichogrammatidae has been unstable. Characteristics that reliably separate females have yet to be found. Male genitalia are diagnostically useful but development of a more reliable classification has been slowed due to the paucity of males. The males have antennae with broadly feathered cilia (Pl. 1, fig. h), whereas the female antennae have fewer and much shorter cilia. The genus Trichogramma is the most commonly used biological control agent of economically important lepidopterous larvae (Li 1994) with the life histories of some species, particularly T. pretiosum, extensively studied. To locate hosts, female wasps primarily rely upon airborne chemical cues associated with scales left near the egg by female moths during oviposition (Beevers et al. 1981), but females also use visual cues such as shape and colour to determine the suitability of a potential host (Ruberson and Kring 1993). While not yet confirmed, in light of recent observations, it is almost certain that T. pretiosum females use chemical cues to avoid intraspecific and interspecific competition over host eggs (Carneiro and Fernandes 2012). Such recognition and restraint minimises the potential for competition between individuals of a single species (super parasitism) and between parasitoid species (multiple parasitism) within a single host. Trichogramma pretiosum undergoes three instars, completes larval development in as little as three days, and during summer, a life cycle is complete in less than 10 days (Strand 1986). During the winter, partially developed larvae remain dormant within the host egg (López and Morrison 1980). Braconidae (Ichneumonoidea) The braconids are another large parasitoid family with more than 40 subfamilies, approximately 1000 genera, 12 000 recognised species, and an estimated 40 000+ species yet to be named (Whitfield et al. 2004). There are records for two subfamilies (Rogadinae and Microgasterinae) associated with Sphingidae in Australia (see Appendix 1). The single rogadine record from Australia is based upon multiple adults of a Macrostomion species emerging from an Acosmeryx anceus anceus (Stoll) larva (Pl. 3, fig. b). Macrostomion species have been reported in association with Gnathothlibus eras Boisduval from Papua New Guinea (Shaw 2002) and Theretra silhetensis (Walker) from Japan (Maeto and Arakaki 2005), sphingid species whose ranges extend into Australia. In both instances, a number of the endoparasitic wasps emerged simultaneously from the mummified prepupal carcass of the host. Microplitis species of the subfamily Microgasterinae are also endoparasitoids. These robust wasps complete their larval development prior to the host reaching larval maturity. Once the host larva reaches the 3rd instar, the parasitoid larva breaks through the cuticle and spins a substantial silken cocoon on the back of the host (Pl. 53, figs e–g). As a result of the relatively small size of the host and large size of the wasp, Microplitis species are solitary parasitoids. In Australia, Microplitis has been documented using Psilogramma argos Moulds and Lane (Pl. 3, fig. a), Cephonodes kingii (W.S. Macleay) and Theretra oldenlandiae (Thon) 190613 Hawkmoths of Australia 3pp.indd 32 (Austin and Dangerfield 1992). Further collecting is expected to result in additional Microplitis-sphingid associations for Australia. Ichneumonidae (Ichneumonoidea) The ichneumonids are the largest parasitoid family with an estimated 60 000+ species worldwide (Gauld 1987). Previously, there were no documented associations between ichneumonids and sphingids in Australia. There are now records for two species and both are associated with the same host, Coequosa triangularis (Appendix 1). Ichneumonids can be either ectoparasitoids or endoparasitoids and are reported to attack lepidopteran larvae and pupae (Townes 1958). Among the larger species, there tends to be one parasitoid per host as the largest and fastest developing wasp larva kills any competitors. In Australia, Lissopimpla excelsa (Costa) (Pimplinae) and Netelia sp. (Tryphoninae) are larval parasitoids. The C. triangularis larvae that hosted them were securely sleeved following their discovery in the midinstars (T. Deane pers. comm.) suggesting the attacks took place early in their development. The Netelia and L. excelsa wasps emerged from the final larval instar of the host (Pl. 3, figs d, e). Gavin Broad (pers. comm.) indicated that L. excelsa is a pupal parasitoid but dissection of the C. triangularis larval carcass yielding the L. excelsa wasp, revealed a large (14mm x 6mm), empty, silken ichneumonid cocoon. Perhaps this is a host-specific L. excelsa adaptation to the unusually large larva and long larval period of C. triangularis which, along with C. australasiae (Donovan), are the only sphingids known to overwinter in the larval stage. Scelionidae (Platygastroidea) There are 58 genera of Scelionidae in Australia representing 763 species (Australian Biological Resources Study 2009). Among the Australian wasp families known to attack sphingid eggs (Appendix 1), scelionids are the most frequently encountered in Australia as well as in North America. To date, only Telenomus in the subfamily Telenominae is known to use sphingid eggs in Australia. Fifty-nine Telenomus species are recognised from Australia (Australian Biological Resources Study 2009) and 9 are now associated with sphingids (Appendix 1). Telenomus species (Pl. 1, fig. f) are recognised as exclusively primary parasitoids; there are no records of them acting as hyperparasitoids. However, some scelionids are subject to hyperparasitoidism by three families of Chalcidoidea (Noyes 2017). A number of Telenomus species have been extensively studied for their potential as biocontrol agents of economic pests in the Americas, Europe and Africa, and several nonnative Telenomus species have been introduced into Australia for that purpose. However, studies have not been undertaken on native Australian Telenomus populations (Don Sands pers. comm.), so very little is known about their biology. Johnson (1988) noted that in Australia a host is recorded for only two species of Telenomus, a pentatomid (Hemiptera) and a saturniid (Lepidoptera). Further notes on the biology of some exotic Telenomus species can be found in Rabb and Bradley (1970), Carneiro and Fernandes (2012), Peñaflor et al. (2012), Bueno et al. (2008), Gerling (1972), Pomari et al. (2012), Legault et al. (2012) and Torgersen and Ryan (1981). There are some biological observations we can report from unidentified Australian Telenomus species. During the summer season in the wet tropics, development of the parasitoid larva takes little more than a week. A day or two before emergence, the developing wasps become clearly discernible through the eggshell (Pl. 23, fig. b). The number of 29/08/19 11:11:07.53 BIOLOGY wasps emerging from a single host egg may vary; some host eggs produce a single wasp, whereas others produce several wasps. This variation does not appear to be a function of egg size or parasitoid size. In contrast, in hundreds of replications, T. remus produces only one wasp per host egg (Bueno et al. 2008; Gerling 1972). Diptera Four dipteran families have been associated with sphingids in Australia. Tachinidae are the most commonly encountered parasitoids of sphingid larvae and play an important role in regulating population levels. Coffin flies (Phoridae) and flesh flies (Sarcophagidae) are often misinterpreted as being parasitoids, but their maggots are scavengers that feed on decaying flesh. In the case of Lepidoptera, they are associated with dead or dying larvae (Pl. 2, fig. f). As previously discussed, adult midges (Ceratopogonidae) are parasites that feed upon the haemolymph of their hosts. Tachinidae There are approximately 10 000 described tachinid species worldwide and thousands of additional species still to be described (Stireman et al. 2006; O’Hara 2008). O’Hara et al. (2004) estimate there are 3500–4000 species found in Australia but less than 500 have been described. Tachinid species (Pl. 2, figs a–e, g, h) are common parasitoids of hawkmoths and Table 1 documents the egg laying strategies of the two subfamilies, six tribes, and 10 genera currently known to be associated with Sphingidae in Australia. In addition to the large number of tachinid-sphingid larval associations documented in the literature, there are many references to tachinid-sphingid pupal parasitoids. However, while some tachinids clearly emerge from the pupae of sphingid hosts (Table 1), we suspect that tachinids are not pupal parasitoids, per se. Depending upon the egg laying strategy of the tachinid parasitoid (Table 1), field collected larvae may show no signs of attack and ‘successfully’ pupate in captivity, only to have tachinids later emerge. In many cases, this has led to the perception that the pupa was attacked. This issue deserves closer examination to determine whether tachinids are exclusively larval parasitoids, but in our extensive rearing, every sphingid pupa giving rise to tachinid adults must have been attacked prior to pupation. Regardless, all tachinids use their hosts as food and shelter for their developing larvae and, as with all parasitoids, such attacks result in the death of the host. Beyond the shared general strategy (host serving as food and shelter for the developing parasitoid larva) and consequence of parasitoidism (death of the host), tachinids use four very distinctive oviposition approaches, three of which have been reported in association with Australian sphingids. The fecundity of females appears related to the probability of successful host infection resulting from each oviposition strategy. Species using a ‘direct’ oviposition strategy (laying eggs directly on, or in, the host) produce far fewer eggs than species using an ‘indirect’ oviposition strategy (laying eggs on the foodplant to be later ingested by the host larva). The first and most common direct oviposition approach used by tachinids attacking sphingids in Australia involves laying translucent fully mature eggs with an extremely thin shell on or very near the host. The first instar parasitoid larva hatches almost immediately after the egg is laid. If the egg is laid on its host, the parasitoid larva immediately burrows into it, but if the egg is laid near the host, the active larva has little difficulty in locating its host. This direct oviposition approach 190613 Hawkmoths of Australia 3pp.indd 33 33 TABLE 1. Egg laying strategies of tachinids currently known to be associated with Sphingidae in Australia. Parasitoid Egg laying strategy Exoristinae Carceliini Carcelia ovolarviparous Exoristini Exorista ovolarviparous Siphonini Ceromya ovolarviparous Sturmiini Blepharipa micro-eggs Drino ovolarviparous Palexorista ovolarviparous Sturmia micro-eggs Zygobothria ovolarviparous Winthemiini Winthemia macro-eggs Tachininae Nemoraeini Nemoraea ovolarviparous is highly successful and females have only a few hundred eggs. This strategy has occasionally been referred to as larviparous but is more correctly referred to as ovolarviparous. The second direct oviposition approach always involves laying large macro-eggs on the cuticle of the host (Pl. 36, fig. h). These eggs are not fully developed and need a few days to mature. In spite of the very sticky adhesive used by the female, we have observed sphingid larvae remove tachinid eggs with their mandibles. The long ovipositor of tachinid females employing this tactic allows them to place their eggs in deep cuticle folds that are difficult for the host larva to reach with its mandibles. Fecundity tends to be slightly higher among females using this strategy. The third direct oviposition approach involves a piercing ovipositor that penetrates the host cuticle and lays eggs internally. Until the current study, there were no documented cases of tachinids using this approach to attack sphingids in Australia, although the genus Compsilura (Exoristinae: Blondeliini), which uses this approach, has been recorded from Queensland and Papua New Guinea (Cantrell and Crosskey 1989). However, recent sampling has documented Compsilura concinnata, which has over 200 insect hosts reported worldwide (Tuttle 2007), attacking Hippotion celerio in Malanda, Queensland. Given the high probability of successful host infection, females using this strategy have very low fecundity levels. The fourth approach involves an indirect oviposition strategy. Micro-eggs (eggs barely visible to the naked eye) are laid on the foodplant and are then ingested by the feeding sphingid larva. The female parasitoids often ‘stalk’ host larvae, patiently wait until feeding commences, and then hurriedly lay their eggs in the freshly cut feeding swathe. Larvae thus infested may be responsible for the perception that some tachinids are pupal parasitoids. Fecundity of tachinids producing micro-eggs is very high, females often contain several thousand eggs. 29/08/19 11:11:07.61 Classification and nomenclature Higher classification Kitching and Cadiou’s (2000) annotated revisionary checklist of the world’s Sphingidae provided a modern and well-founded higher classification that we follow in this book. These authors presented a lengthy discussion on the evolving ideas about the higher classification. They emphasised the significant contribution made by Rothschild and Jordan (1903) in formulating ideas that set a sound foundation for modern sphingid classification. Since Rothschild and Jordan, a number of alternative classifications have been proposed (Janse 1932; Carcasson 1968; Hodges 1971; Nakamura 1976, 1977, 1978; Derzhavets 1984; Minet 1994) and carefully evaluated by Kitching and Cadiou. Adopting the principles of the classification proposed by Minet (1994), Kitching and Cadiou (2000) recognised three subfamilies, the Smerinthinae, Sphinginae and Macroglossinae, all of which occur in Australia. Within their descending hierarchy of classification, Kitching and Cadiou recognised three tribes within the Smerinthinae, all found in Australia, two tribes within the Sphinginae also found in Australia, and three tribes and four subtribes within the Macroglossinae, all of which occur in Australia except the small tribe Philampelini and the large subtribe Dilophonotina. Kitching and Cadiou’s classification is based primarily upon unpublished cladistic analyses by Kitching but they emphasise that their classification is provisional, still under study, and that the placement of some genera should be considered tentative. In particular, they refer to the placement of four Australian genera, Synoecha Rothschild and Jordan, Coenotes Rothschild and Jordan, Hopliocnema Rothschild and Jordan and Tetrachroa Rothschild and Jordan. Kitching and Cadiou transferred Coenotes from the Sphingini to the Sphingulini to be placed closest to Synoecha. However, they go on to say that the inclusion of all four genera within the Sphingulini remains to be confirmed. Kamaluddin et al. (2014) attempted an intuitive phylogeny based on morphological characters for a wide selection of genera found in Pakistan and Azad Kashmir, and while this approach has limitations, it provides some interesting attributes at nodes, not unlike the trees provided by Rothschild and Jordan (1903). Recent molecular phylogenetic studies (Regier et al. 2001; Hundsdoerfer, Kitching and Wink 2005; Kawahara et al. 2009) for the most part concur with the conclusions of Kitching and Cadiou (2000) although some disparities have been highlighted. The most comprehensive of these studies, and the most recent, are those of Kawahara et al. (2009), which included 131 species, and of Kawahara and Barber (2015), which included an additional 84 species, both studies representing all currently recognised subfamilies, tribes and subtribes. While their results provided strong support for groups such as the Macroglossinae, Sphinginae, Acherontiini, Ambulycini, Choerocampina and Hemarina, they found some groups were paraphyletic or polyphyletic, e.g. that the Smerinthinae and the tribes Dilophonotini, Macroglossini and Smerinthini were paraphyletic with respect to Sphinginae. Among other discrepancies, they also found that the Acherontiini (sister to which was a group of genera centred around Psilogramma) fell within the Sphingini rendering the 190613 Hawkmoths of Australia 3pp.indd 34 Sphingini paraphyletic, and they suggested a solution would be to confer tribal status to the Psilogramma group. Further, they found that species within the Macroglossinae differed in relationships in several ways from the generally accepted classification within the subfamily although support for their nodes was not particularly strong. Kawahara et al. (2009) and Kawahara and Barber (2015) refrained from making formal changes to the higher classification pending more comprehensive studies although some have been recently made by Kitching et al. (2018a). Primarily, Kitching et al. moved the tribe Sphingulini from the subfamily Smerinthinae to the Sphinginae and suggested the likely need for two new subtribes in the Macroglossini, three new tribes in the Smerinthinae, a new tribe in the Sphinginae, and a new subtribe in the Sphingini but formal designations were not made pending further study. Zolotuhin and Ryabov (2012) proposed an alternative arrangement for some tribes and resurrected some old names from Hübner (1819) in a tentative re-classification pointing out the uncertainties around the higher classification of the Sphingidae. They proposed the name Bombyliinae replace Macroglossinae and the replacement of some tribal names in current use. We use the names established by Kitching and Cadiou (2000) and amended by Kitching et al. (2018b), pending a stable higher classification, to avoid confusion in the interim. Genus and species There have also been extensive molecular studies at the species level. Most analyses have been done in the Barcode of Life Project (BOLD), which has the Sphingidae as one of its target groups. Many of the world species have been sequenced for the COI gene in an attempt to provide a unique barcode for identifying each species. The Australian fauna has been particularly well documented. Many of these sequences are in the public domain and are being used by taxonomists to find cryptic species and in sorting difficult species complexes, although only few published papers have so far acknowledged the use of BOLD data. Since the publication by Kitching and Cadiou (2000), we recognise seven generic changes within the Australian fauna. Five new genera were created including: Zacria Haxaire and Melichar to accommodate a newly discovered species, Z. vojtechi (Haxaire and Melichar 2003); Cerberonoton Zolotuhin and Ryabov for C. rubescens severina (now C. severina), which was removed from Meganoton Boisduval (Zolotuhin and Ryabov 2012); Imber Moulds, Tuttle and Lane for the Australian endemic I. tropicus, which was removed from Langia Moore (Moulds, Tuttle and Lane 2010); Pseudoangonyx Eitschberger for P. excellens, which was removed from Angonyx (Eitschberger 2010c); and Chelacnema for C. ochra, which we remove from Hopliocnema in this book. At the species level, the number of named sphingid species has increased dramatically since Kitching and Cadiou (2000). The genus most impacted was Psilogramma which increased from five to 61 species before Kitching et al. (2018b) synonymized many of these names, reducing the number to 29 species. 29/08/19 11:11:07.68 CLASSIFICATION AND NOMENCLATURE Among the Australian fauna, Kitching and Cadiou (2000) recognised only two Psilogramma species (one listed only in the Appendix, added in press). Yet within the next year, three notable reviews of the genus Psilogramma were published (Brechlin 2001; Eitschberger 2001a, 2001b) that raised the number of described Australian Psilogramma species to six. Subsequent papers (Eitschberger 2004a, 2010a, 2010b; Brechlin and Kitching 2010b; Lane, Moulds and Tuttle 2011) increased that number to ten. Three of these species were later synonymised by Eitschberger (2010b), Brechlin and Kitching (2010b) and Rougerie et al. (2014) leaving the seven species currently recognised, viz. P. casuarinae (Walker, 1856), removed from synonymy with P. menephron; P. menephron (Cramer, 1780); P. argos Moulds and Lane, 1999; P. papuensis Brechlin, 2001; P. maxmouldsi Eitschberger, 2001; P. exigua Brechlin, Lane and Kitching, 2010; and P. penumbra Lane, Moulds and Tuttle, 2011. One additional species, P. discistriga discistriga, is found on Christmas Island. Resolving the identity of the Australia Psilogramma species has been a complex process and is discussed in detail in the genus and relevant species treatments. In addition to Psilogramma species, other Australian species described since Kitching and Cadiou (2000) include Zacria vojtechi (Haxaire and Melichar 2003), Gnathothlibus 190613 Hawkmoths of Australia 3pp.indd 35 35 australiensis (Lachlan 2004b), Hopliocnema lacunosa and H. ochra (Tuttle, Moulds and Lane 2012), and Coenotes arida (Moulds and Melichar [2014]). Further, Macroglossum corythus approximans (as M. stenoxanthum) was removed from synonymy by Eitschberger (2003d), Theretra celata was given species status (Vaglia et al. 2010) and, in this book, Hippotion johanna (Kirby) is removed from synonymy with Hippotion brennus (Stoll), Acosmeryx cinnamomea (Herrich-Schäffer) is removed from synonymy with Acosmeryx anceus anceus (Stoll), and Cephonodes australis, C. cunninghami, Cerberonoton severina, Macroglossum errans, M. papuanum and M. queenslandi are all elevated to species status. Species names deleted from the Australian fauna in this book are: Cephonodes hylas (Linnaeus, 1771), C. picus (Cramer, 1777), Cerberonoton rubescens (Butler, 1876), Macroglossum hirundo (Boisduval, 1832), Macroglossum troglodytus Boisduval, [1875] and Macroglossum heliophila Boisduval, [1875]. Species not previously recorded from Australia are Marumba timora from the Kimberley coast, Amplypterus panopus panopus from Darwin, Macroglossum melas melas and Cypa decolor euroa from the Torres Strait Islands and Daphnis hypothous crameri, Macroglossum ungues ungues, Psilogramma discistriga discistriga and Theretra lucasii from Christmas Island. 29/08/19 11:11:07.73 The Australian fauna Checklist of Australian species Subfamily MACROGLOSSINAE Harris, 1839 The Australian sphingid fauna currently comprises 87 species (89 taxa) in 31 genera. Twenty-four species are recognised at subspecies level. Higher classification and nomenclature follow Kitching and Cadiou (2000), Kawahara et al. (2009) and Kitching et al. (2018a, 2018b). Subsequent changes to genus and species nomenclature are discussed in detail in the relevant genera and species treatments. Tribe Hemarini Tutt, 1902 Cephonodes australis Kitching and Cadiou, 2000 stat. nov. Cephonodes cunninghami (Cramer, 1777) stat. nov. Cephonodes janus Miskin, 1891 Cephonodes kingii (W.S. Macleay, 1826) Superfamily SPHINGOIDEA Latreille, [1802] Tribe Macroglossini Harris, 1839 Subtribe Macroglossina Harris, 1839 Angonyx papuana papuana Rothschild and Jordan, 1903 Cizara ardeniae (Lewin, 1805) Daphnis dohertyi dohertyi Clark, 1922 Daphnis hypothous crameri Eitschberger and Melichar, 2010 Daphnis moorei (W.J. Macleay, 1866) Daphnis placida placida (Walker, 1856) Daphnis protrudens protrudens C. and R. Felder, 1874 Eupanacra splendens splendens (Rothschild, 1894) Eurypteryx molucca C. and R. Felder, 1874 Gnathothlibus australiensis Lachlan, 2004 Gnathothlibus eras (Boisduval, 1832) Macroglossum alcedo Boisduval, 1832 Macroglossum corythus corythus Walker, 1856 Macroglossum corythus approximans T.P. Lucas, 1891 stat. nov. Macroglossum dohertyi doddi Clark, 1922 Macroglossum errans Walker, 1856 stat. nov. Macroglossum joannisi Rothschild and Jordan, 1903 Macroglossum melas melas Rothschild and Jordan, 1903 Macroglossum micacea micacea Walker, 1856 Macroglossum nubilum Rothschild and Jordan, 1903 Macroglossum papuanum Rothschild and Jordan, 1903 stat. nov. Macroglossum prometheus lineata T.P. Lucas, 1891 Macroglossum prometheus prometheus Boisduval, [1875] Macroglossum queenslandi Clark, 1927 stat. nov. Macroglossum rectans Rothschild and Jordan, 1903 Macroglossum tenebrosa T.P. Lucas, 1891 Macroglossum ungues ungues Rothschild and Jordan, 1903 Macroglossum vacillans Walker, [1865] Nephele hespera (Fabricius, 1775) Nephele subvaria (Walker, 1856) Pseudoangonyx excellens (Rothschild, 1911) Zacria vojtechi Haxaire and Melichar, 2003 Subtribe Acosmerygina Tutt, 1904 Acosmeryx anceus anceus (Stoll, 1781) Acosmeryx cinnamomea (Herrich-Schäffer, [1869]) stat. rev. Acosmeryx miskini (Murray, 1873) Family SPHINGIDAE Latreille, [1802] Subfamily SMERINTHINAE Grote and Robinson, 1865 Tribe Ambulycini Butler, 1876 Ambulyx wildei Miskin, 1891 Ambulyx dohertyi queenslandi Clark, 1928 Amplypterus panopus panopus (Cramer, 1779) Tribe Smerinthini Grote and Robinson, 1865 Imber tropicus (Moulds, 1983) Coequosa australasiae (Donovan, 1805) Coequosa triangularis (Donovan, 1805) Cypa decolor euroa Rothschild and Jordan, 1903 Tribe Sichiini Tutt, 1902 Marumba timora Rothschild and Jordan, 1903 Subfamily SPHINGINAE Latreille, [1802] Tribe Sphingulini Rothschild and Jordan, 1903 Chelacnema ochra (Tuttle, Moulds and Lane, 2012) comb. nov. Coenotes arida Moulds and Melichar, [2014] Coenotes eremophilae (T.P. Lucas, 1891) Hopliocnema brachycera (Lower, 1897) Hopliocnema lacunosa Tuttle, Moulds and Lane, 2012 Synoecha marmorata (T.P. Lucas, 1891) Tetrachroa edwardsi (Olliff, 1890) Tribe Sphingini Latreille, [1802] Subtribe Sphingina Latreille, [1802] Cerberonoton severina (Miskin, 1891) stat. rev. Leucomonia bethia (Kirby, 1877) Psilogramma argos Moulds and Lane, 1999 Psilogramma casuarinae (Walker, 1856) Psilogramma discistriga discistriga (Walker, 1856) Psilogramma exigua Brechlin, Lane and Kitching, 2010 Psilogramma maxmouldsi Eitschberger, 2001 Psilogramma menephron nebulosa (Butler, 1876) Psilogramma papuensis Brechlin, 2001 Psilogramma penumbra Lane, Moulds and Tuttle, 2011 Subtribe Acherontiina Boisduval, [1875] Acherontia lachesis (Fabricius, 1798) Agrius convolvuli (Linnaeus, 1758) Agrius godarti (W.S. Macleay, 1826) Megacorma obliqua obliqua (Walker, 1856) 190613 Hawkmoths of Australia 3pp.indd 36 29/08/19 11:11:07.81 THE AUSTRALIAN FAUNA Subtribe Choerocampina Grote and Robinson, 1865 Hippotion boerhaviae (Fabricius, 1775) Hippotion brennus (Stoll, 1782) Hippotion celerio (Linnaeus, 1758) Hippotion johanna (Kirby, 1877) stat. rev. Hippotion rosetta (Swinhoe, 1892) Hippotion scrofa (Boisduval, 1832) Hippotion velox (Fabricius, 1793) Hyles livornicoides (T.P. Lucas, 1892) Theretra celata celata (Butler, 1877) Theretra indistincta indistincta (Butler, 1877) Theretra inornata (Walker, [1865]) Theretra insularis insularis (Swinhoe, 1892) Theretra latreillii (W.S. Macleay, 1826) Theretra lucasii (Walker, 1856) stat. rev. Theretra margarita (Kirby, 1877) Theretra nessus nessus (Drury, 1773) Theretra oldenlandiae (Thon, 1828) Theretra queenslandi (T.P. Lucas, 1891) Theretra silhetensis intersecta (Butler, [1876]) Theretra tryoni (Miskin, 1891) Theretra turneri (T.P. Lucas, 1891) Key to last instar larvae The last instar larvae of many sphingid species can be extremely variable in both colour and markings and new variations continue to be discovered. As a result, it has not been possible for this key to address all of the variants that we have encountered, let alone anticipate the many yet to be discovered. This is especially true with respect to some members of the genera Macroglossum and Cephonodes. The key includes 71 species and is based on examination of living larvae supplemented by colour photographs. Supplementary information is often given in brackets [ ] to help provide clarity in some couplets. In situations where the larval foodplant has been identified, it also may be helpful to consider Appendix 2, ‘Summary of known larval foodplants’. Larval associations with plant species, genera or even families may provide valuable leads towards identifications. As an example, without context, a variant of Macroglossum errans may be difficult to distinguish from some individuals of M. vacillans or M. micacea micacea; however, to date in Australia, M. errans is only known to feed on species of the family Rubiaceae and M. vacillans and M. micacea micacea only on species of the genus Strychnos in the family Loganiaceae. 1 – 2 – 3 – 4 – Caudal horn lacking ...................................................... 2 Caudal horn always present although the size and form varies greatly (Figs 44–46) ................................. 4 Small larva not exceeding 50 mm in length (Fig. 46) .. ................................... (Pl. 34) Hopliocnema brachycera Large larva 95–125 mm in length ................................ 3 Claspers with a conspicuous, sclerotised, black, glossy, bead-like ‘false eye’ ..... (Pl. 20) Coequosa triangularis Claspers without a ‘false eye’............................................ ....................................... (Pl. 19) Coequosa australasiae Prominent circular to elliptical subdorsal eyespot(s) present (Fig. 44) [always only one pair per segment, one visible on each side; some eye-like in appearance with a ‘pupil’ and/or interior dots] [not to be confused with a lateral patch surrounding a spiracle, or a coloured patch within a lateral stripe] ........................ 5 Eyespot(s) absent .......................................................... 29 190613 Hawkmoths of Australia 3pp.indd 37 5 – 6 – 7 – 8 – 9 – 10 – 11 – 12 – 13 – 14 – 15 – 16 – 17 – 18 – 19 37 Eyespot(s) present on the abdominal segment(s) and/ or thoracic segment(s) [use caution when examining photographs, as it may be diﬃcult to interpret the eyespot’s placement; if the eyespot is above a spiracle, it is abdominal] ............................................................... 6 Eyespot(s) present only on abdominal segment(s) .... 10 Eyespot present only on the metathoracic segment ..... ............................................................................................ 7 Eyespots present on the thoracic and abdominal segments .......................................................................... 8 Spiracles solid dark reddish orange to nearly black .... ................................................... (Pl. 21) Daphnis moorei Spiracles yellowish white to pale yellow, vertically bisected by a thin black line ............................................. ...................... (Pl. 23) Daphnis protrudens protrudens Eyespots boldly highlighted with black [in some specimens, lower half of the eyespots obscured by a bold, intersecting whitish/yellowish subdorsal stripe] .............................................. (Pl. 35) Hyles livornicoides Eyespots lack black highlighting ................................. 9 Eyespot(s) faintly and very narrowly encircled ............. ............................. (Pl. 25) Gnathothlibus australiensis Eyespot(s) encircled with a broad, green to bluish green ‘halo’, particularly pronounced on the ﬁrst few abdominal segments ....... (Pl. 26) Gnathothlibus eras Eyespot only on abdominal segment 1 ..................... 11 Eyespot on abdominal segment 1 and one or more additional abdominal segments ................................. 14 Eyespot without red pigmentation in the ‘pupil’ .... 12 Eyespot with bold red pigmentation in the ‘pupil’ ....... ................................................. (Pl. 64) Theretra latreillii Eyespot lacks a white or pale blue ‘pupil’ and does not bulge convexly above the abdominal segment (surface smooth) [caudal horn tapers, never bluntly ﬂattened] ... .......................................................................................... 13 Eyespot has a prominent white or pale blue ‘pupil’ and convexly bulges above the abdominal segment (subtle in some individuals) [stout caudal horn with a bluntly ﬂattened black tip]........ (Pl. 68) Theretra queenslandi Caudal horn long, usually thin, often whip-like and curving forward .................... ( Pl. 33) Hippotion velox Caudal horn short, stout and curving backwards ........ ...................... (Pl. 24) Eupanacra splendens splendens Eyespots only on abdominal segments 1 and 2 ........ 15 Eyespots on abdominal segments 1 and 2 plus others, usually segments 1–7 and occasionally 1–8 ............... 19 Eyespots on abdominal segments 1 and 2 the same in size and markings ......................................................... 16 Eyespots on abdominal segments 1 and 2 vary in size and/or markings ............................................................ 17 Caudal horn concolourous tending pinkish red (lacking a pale distal portion) .......................................... ................................................ (Pl. 63) Theretra inornata Caudal horn with a distinct oﬀ-white to pale dull yellow distal tip......................... (Pl. 70) Theretra tryoni Eyespots on abdominal segments 1 and 2 are similar in markings, varying in size only ............................... 18 Eyespots on abdominal segments 1 and 2 are not similar in markings or size ............................................... ................................................. (Pl. 29) Hippotion celerio Eyespots lacking a ‘pupil’, caudal horn tapering to a slender point without a black tip ..................................... ....................................... (Pl. 66) Theretra nessus nessus Eyespots with a multicoloured yellow, green and blue ‘pupil’ [caudal horn stout with a bluntly ﬂattened black tip] ........................ (Pl. 68) Theretra queenslandi Seven (7) eyespots present .......................................... 20 29/08/19 11:11:07.90 38 HAWKMOTHS OF AUSTRALIA Figs 44–46. Diagrammatic representations of hawkmoth larvae illustrating morphological features mentioned in the key. – 20 – 21 – 22 – 23 – Eight (8) eyespots present ........................................... 28 Eyespots on all abdominal segments similar in markings ........................................................................ 21 Eyespots on abdominal segments vary in markings, most notable diﬀerence between eyespot on abdominal segment 1 and subsequent eyespots .......................... 25 Eyespot on segment 1 stippled randomly with small, bright white dots, ‘pupil’ not apparent [caudal horn long, thin, straight with a prominent white tip] ........... ........................................ (Pl. 27) Hippotion boerhaviae Eyespot on segment 1 lacking such dots, although in some species a single oﬀ-white to bluish dot may centre a well-deﬁned ‘pupil’ ........................................ 22 Caudal horn sharply hooked backwards, long and often greatly exceeding 6.0 mm .................................. 23 Caudal horn straight, short and always less than 4.0 mm ............................................................................ 24 Eyespots with a semi-circular ‘pupil’ oﬀ-set in the top half of all eyespots ....... (Pl. 61) Theretra celata celata Eyespots with a circular ‘pupil’ centred in all eyespots ....................... (Pl. 62) Theretra indistincta indistincta 190613 Hawkmoths of Australia 3pp.indd 38 24 – 25 – 26 – Tumidity at base of caudal horn not developed so that the junction between the anal plate and abdominal segment 8 when viewed laterally is straight or nearly so (see text-ﬁgures 107a, 108b) [caudal horn less than 2 mm long]....................... (Pl. 65) Theretra margarita Tumidity at base of caudal horn large and conical so that the junction between the anal plate and abdominal segment 8 when viewed laterally is almost 90° (see text-ﬁgures 107b, 108a) [caudal horn more than 3mm long] .................................................................. .......................... (Pl. 69) Theretra silhetensis intersecta Eyespot on abdominal segment 2 very diﬀerent in size and appearance from any other eyespot......................... .................................................. (Pl. 32) Hippotion scrofa Eyespot on abdominal segment 2 similar to at least one additional eyespot ................................................. 26 Caudal horn almost absent, pimple-like, virtually without a shaft, not exceeding 1 mm ............................. .............................................. (Pl. 28) Hippotion brennus Caudal horn clearly developed with a shaft, at least 3 mm long ...................................................................... 27 29/08/19 11:11:08.36 THE AUSTRALIAN FAUNA 27 Eyespots on abdominal segments 5–7 (three most posterior eyespots) with the dominating pale area very elongate and tending parallel-sided ....................... ............................................. (Pl. 30) Hippotion johanna – Eyespots on abdominal segments 5–7 with the dominating pale area tending circular, upper margin almost straight but lower portion rounded ................... ................................................ (Pl. 31) Hippotion rosetta 28(19) Eyespots on abdominal segments 1 and 2 similar, but diﬀerent from all others [caudal horn long, thin, black with a prominent white tip] .............................................. ........................................ (Pl. 67) Theretra oldenlandiae – Eyespots on all segments similar [caudal horn long, thin, entirely black] ................ (Pl. 71) Theretra turneri 29(4) Linear lateral abdominal markings present across several segments (disregard longitudinal middorsal lines), may be vertical, oblique and/or longitudinal (subdorsal, and/or subspiracular) (Fig. 44); although one or a combination is always present, may vary from well-deﬁned and intact to very faint, thin and broken to the point of being little more than short dashes ........ 30 – Linear lateral abdominal markings lacking; abdominal segments a mosaic of brown, grey, green and white, densely stippled with small whitish dots [a pair of large green to brown elliptical to quadrantshaped markings radiating downward from middorsum at the posterior edge of each segment] ...... .......................... (Pl. 40) Macroglossum dohertyi doddi 30 Linear abdominal markings present as vertical bands; abdominal segments with bold pinkish intersegmental bands, ground colour uniformly pale green; spiracles blue and vertically bisected by a thin black line, encircled in yellow; prothoracic shield, anal plate and anal claspers with prominent pinkish red tubercles tipped in white [two forms; most with a prominent subdorsal stripe; the second lacking such a stripe]....... ..................................................... (Pl. 36) Imber tropicus – Linear abdominal markings present as either oblique and/or longitudinal stripe(s) ....................................... 31 31 Only one type of abdominal stripe(s) present, either oblique or longitudinal ................................................ 32 – Both oblique and longitudinal abdominal stripes (may be faint) present in the same individual .................... 42 32 Only oblique abdominal stripe(s) present ................ 33 – Only longitudinal abdominal stripe(s) present ........ 49 33 The oblique abdominal stripes originate at or near the anterior edge of the abdominal segment, continue just above that segment’s spiracle and terminates in the upper subdorsal area of the next segment ................ 34 – The oblique abdominal stripes originate at the spiracle of the abdominal segment and terminate on the dorsum of the next segment, merging into a middorsal stripe [the caudal horn is blue with many yellowish tubercles] ....................................... (Pl. 14) Cerberonoton severina 34 Anal plate with very tiny tubercles (i.e. diﬃcult to see with the naked eye) or lacks tubercles (smooth) ...... 35 – Anal plate with many large tubercles present (clearly visible with the naked eye) .......................................... 38 35 Caudal horn strongly curled forward at the apex, almost forming a ring (text-ﬁgure 66) ........................... .......................................................... Acherontia lachesis – Caudal horn curved backwards in a sweeping arc ....... .......................................................................................... 36 36 Spiracles with a vertically elliptical black centre encircled by orange, blue or white (not always 190613 Hawkmoths of Australia 3pp.indd 39 39 conclusive beyond Australia) ........................................... .................................................. (Pl. 7) Agrius convolvuli – Spiracles unicolourous ................................................. 37 37 Spiracles orange or white [colour of the spiracle contrasts sharply with the ground colour]..................... ........................................................ (Pl. 8) Agrius godarti – Spiracles light greenish with a faint purple tinge [colour of spiracles closely matches the ground colour] ........................................... (Pl. 60) Tetrachroa edwardsi 38 Anal plate with large tubercles much larger than those on caudal horn, clearly larger than any others on larva (2 species, indistinguishable) [individuals may or may not have large purplish brown markings] ...................... (Pls 57, 58) Psilogramma menephron nebulosa or P. papuensis – Anal plate with largest tubercles similar in size or smaller than those on caudal horn, no larger than any others on larva ............................................................... 39 39 Meso- and metathorax with numerous small white tubercles of similar size in many transverse rows (2 species, indistinguishable) [individuals of both species may or may not have large purplish brown markings] ..... (Pls 54, 55) Psilogramma casuarinae or P. exigua – Meso- and metathorax with about 12 large non-white tubercles diﬀering slightly in size in 3–4 transverse rows ................................................................................ 40 40 Meso- and metathoracic tubercles predominantly black ................................... (Pl. 37) Leucomonia bethia – Meso- and metathoracic tubercles never black, predominantly yellow or yellow capped pinkish orange ............................................................................. 41 41 All meso- and metathoracic tubercles concolourous pale yellow [individuals may or may not have large purplish brown body markings] ...................................... ................................ (Pl. 56) Psilogramma maxmouldsi – Some meso- and metathoracic tubercles capped pinkish orange [individuals may or may not have large purplish brown body markings] ...................................... ............................................. (Pl. 53) Psilogramma argos 42(31) Oblique abdominal stripes include two bold white slashes, one transversing abdominal segments 2 and 3 and one on abdominal segment 7 angled in the opposite direction ......................................................... 43 – Oblique abdominal stripes end at the subdorsal stripe and are all angled in the same direction ................... 44 43 Caudal horn 5.5–6.0 mm long [widespread across much of northern Australia] ............................................ ................................................. (Pl. 52) Nephele subvaria – Caudal horn 7.5–8.5 mm long [northern Cape York Peninsula and Torres Strait Islands only] ...................... ................................................... (Pl. 51) Nephele hespera 44 Oblique abdominal stripes angled forwards from their base (toward the larval head) ........................................... ................................................... (Pl. 16) Cizara ardeniae – Oblique abdominal stripes angled backwards from their base (away from the larval head) ...................... 45 45 Small larva (40–55 mm), caudal horn with a white to pale yellowish apical tip ............................................... 46 – Large larva never less than 70 mm long, in some species reaching 105 mm, caudal horn hooked backwards, usually, but not always, with a black apical tip. ................................................................................... 47 46 Caudal horn (4–5 mm), straight ...................................... ................................. (Pl. 45) Macroglossum papuanum – Caudal horn (7–8 mm), apical half curved forward in a shallow arc ............. (Pl. 44) Macroglossum nubilum 29/08/19 11:11:08.44 40 HAWKMOTHS OF AUSTRALIA 47 Densely and boldly stippled with small white to yellowish dots ................................................................ 48 – Lacks stippling or very faintly stippled with small pale yellow or white dots ........................................................... ...................................... (Pl. 6) Acosmeryx cinnamomea 48 Longitudinal pale yellow to white subdorsal stripe; broad stripe bordered above by a narrow dark reddish brown to black stripe ......................................................... ................................... (Pl. 4) Acosmeryx anceus anceus – Longitudinal pale yellow to white subdorsal stripe; broad stripe bordered below by a broad pink stripe and above by a narrow dark green stripe ....................... ................................................ (Pl. 6) Acosmeryx miskini 49(32) Prominent prothoracic shield densely stippled with small white or pale yellow tubercles; spiracles entirely white or white bisected by a reddish orange transverse band ................................................................................ 50 – Prothoracic shield not as described above; spiracles not as described above ................................................. 53 50 Spiracles entirely white ................................................ 51 – Spiracles white but horizontally bisected by a broad reddish orange band ..................................................... 52 51 Caudal horn stout, hooked backwards [two forms, ﬁrst with entire ground colour a concolourous light green ground colour, second with green ground colour but the region between the subdorsal and subspiracular stripe tinged orange]; spiracles boldly encircled with orange, black splotches often present (highly variable) ................. (Pl. 12) Cephonodes janus – Caudal horn thin, and of a shallow ‘S’ shape (highly variable) ............................... (Pl. 13) Cephonodes kingii 52 Larva with a glossy, almost oily, appearance; prothoracic shield with anterior row of tubercles similar in size to subsequent rows (highly variable) .... ................................. (Pl. 11) Cephonodes cunninghami – Larva with a matt appearance; prothoracic shield with anterior row of tubercles distinctly larger than subsequent rows (highly variable) ................................... ......................................... (Pl. 10) Cephonodes australis 53 Spiracles embedded within an enlarged, often irregularly shaped blotch (not to be confused with a thin band encompassing the spiracles) ..................... 54 – Spiracles not embedded within a blotch (if in doubt, treat as not in such a blotch) ........................................ 59 54 Blotches surrounding the spiracles grey, blackish, greyish purple or green ................................................ 55 – Blotches surrounding the spiracles are red, orange or orangish brown ............................................................. 56 55 Spiracles orange ....................... (Pl. 72) Zacria vojtechi – Spiracles black..................................................................... ............ (Pl. 39) Macroglossum corythus approximans 56 Larval face has a pair of vertical stripes .................... 57 – Larval face lacks vertical stripes and is generally unmarked ....................................................................... 58 57 A prominent white subspiracular stripe present, bordered on each side by a ﬁne black line ...................... ..................................................... (Pl. 17) Coenotes arida – A prominent white subdorsal strip lacking (highly variable) ............................................................................... ................. (Pl. 46) Macroglossum prometheus lineata 58 Ground colour black to light grey; a prominent red to gold middorsal stripe, often broken; a subdorsal stripe, white or yellow, is extremely variable from prominent to very faint and broken to lacking; a thin subspiracular stripe (white to yellow) is also present; the caudal horn 190613 Hawkmoths of Australia 3pp.indd 40 is thin and arcs slightly backwards (highly variable) ... ........................................ (Pl. 18) Coenotes eremophilae – Ground colour of dorsum a uniform reddish orange or green (no middorsal stripe) [subdorsal stripe is little more than a transitional line between the boldly coloured dorsum and the contrastingly coloured lateral region], lateral region densely stippled with large white dots] (highly variable) ................................... ...................... (Pl. 43) Macroglossum micacea micacea 59(53) Caudal horn hooked sharply backwards .................. 60 – Caudal horn not hooked sharply backwards ........... 61 60 Caudal horn orange to orangish yellow with a prominent black tip, spiracles solid orange [bluish spots encircled with black are tightly compacted to the point of forming an almost unbroken longitudinal line just below the subdorsal stripe] ................................ ................................... (Pl. 22) Daphnis placida placida – Caudal horn unicolourous orange to orangish yellow, yellowish brown spiracles vertically bisected by a black line ..... (Pl. 23) Daphnis protrudens protrudens 61 Longitudinal abdominal stripes present only above the spiracles (may be very faint, thin and interrupted, little more than dashes) ............................................... 62 – Longitudinal abdominal stripes present both above and below the spiracles in the same individual (may be very faint, thin and interrupted, little more than dashes) ............................................................................ 73 62 One or two abdominal stripe(s) present above the spiracles (may vary from bold, well-deﬁned, and intact to very faint, thin, and broken to the point of being little more than short dashes) .......................... 63 – Three abdominal stripes present above the spiracles [white, yellowish or brownish ground colour with three bold subdorsal stripes, brown to black; a similar stripe is interrupted on each abdominal segment by the spiracle; black caudal horn thin and whip-like] ..... ................................... (Pl. 49) Macroglossum tenebrosa 63 Lateral stripe touching or almost touching the top of the spiracles always present ......................................... 64 – Lateral stripe never close to touching the top of the spiracles .......................................................................... 66 64 A lateral and subdorsal stripe present [lateral stripe bold and bright white, subdorsal stripe thin, faint pale yellowish green, base of the caudal horn on abdominal segment 8 with a bold white slash] .................................. ......................................... (Pl. 59) Synoecha marmorata – Only a lateral stripe present ........................................ 65 65 Spiracles orange, a black horn up to 3.5 mm, atop a prominent red tumidity..... (Pl. 15) Chelacnema ochra – Spiracles black, a very short black horn, never more than just over 1 mm ................ (Pl. 34) Hopliocnema lacunosa 66 Face with vertical stripes (may be faint but always present) ........................................................................... 67 – Face without vertical stripes, generally unmarked....... .......................................................................................... 68 67 Facial stripes thin, white to pale yellow in pale or green forms but in brown forms may be broadly dark brown, [spiracles predominantly orange; caudal horn long and gently sweeps in a forward arc, usually with a pale tip] (highly variable) .................................................. ................. (Pl. 46) Macroglossum prometheus lineata – Facial stripes broad, dark brown [spiracles black, vertically bisected by white; caudal horn long, straight or nearly so, concolourous] .............................................. ................................ (Pl. 9) Angonyx papuana papuana 29/08/19 11:11:08.52 THE AUSTRALIAN FAUNA 68 Ventral portion of thoracic segments with a bold, broad yellow to orangish slash (highly variable); subdorsal stripe may be bold on all abdominal segments or only faint on the more distal segments; stripe may be concolourous or bi-coloured; spiracles orange in green form, dark brown in brown form) ...... ...................... (Pl. 43) Macroglossum micacea micacea – Ventral portion of thoracic segments lack such a slash .......................................................................................... 69 69 Subdorsal stripe is thin and faint but transitions to a broad white slash on abdominal segments 7 and 8, ending at the base of the caudal horn ........................ 70 – Subdorsal stripe lacks such a transition and contrasting slash ........................................................... 71 70 Spiracles entirely orange [caudal horn dark brown to black, gently tapering in a shallow forward arc] (highly variable) ......................... (Pl. 41) Macroglossum errans – Spiracles grey or black, vertically bisected by a white line [caudal horn straight] ................................................ ...................................... (Pl. 42) Macroglossum joannisi 71 Subdorsal stripe is concolourous white to yellow (disregard subtle changes in hue) ............................... 72 – Subdorsal stripe is multicoloured, upper portion is white to yellow and the lower portion is a suﬀused brown or purple of varying shades [purple is greatly reduced in the brown morph] ......................................... .................................... (Pl. 50) Macroglossum vacillans 72 Caudal horn straight, short, 3.5–5.0 mm (two forms, ﬁrst with a mosaic of contrasting shades of brown and grey; a faint yellowish brown subdorsal stripe; a pair of small slightly elongated black spots are just posterior and equidistance above and below the spiracle on each abdominal segment; in brown form, the lower spot may not be visible; a similar spot may or may not be present in the subdorsal region of some individuals; in a second form the above-mentioned spots sometimes lacking; ground colour, except for the bold yellow subdorsal stripe, otherwise an unmarked light green). (Pl. 38) Macroglossum alcedo – Caudal horn a shallow ‘S’ shape, long 7.0–8.9 mm, the apical 1/4 to 1/3 in most individuals white, most then subsequently tipped black [lateral and ventral region a light green, dorsal and subdorsal region a dull white, faintly tinged with light purple. Brown form caudal horn pinkish brown, turning white along its length and apically tipped black .................................................. .............................. (Pl. 47) Macroglossum queenslandi 73(61) Subdorsal stripe usually well-deﬁned; subspiracular stripe evident but often interrupted; abdomen densely stippled with white spots, often of contrasting size, the largest spots most frequently in loose rows of 3–6 on each segment, just below the subdorsal stripe and usually encircled with black ........................................ 74 – Subdorsal stripe very faint, thin and broken, little more than dashes (most evident on posterior abdominal segments); subspiracular stripe only slightly more pronounced; abdomen densely stippled with small white spots but enlarged spots never present .......................... (Pl. 48) Macroglossum rectans 74 Subdorsal and subspiracular stripes the same, or nearly the same, colour ................................................ 75 – Subdorsal and subspiracular stripes not the same colour [subdorsal stripe is multicoloured, upper portion is white to yellow and the lower portion is a suﬀused purple of varying hues (less obvious in the 190613 Hawkmoths of Australia 3pp.indd 41 75 – 41 brown morph), subspiracular stripe is yellowish (form) ........................ (Pl. 50) Macroglossum vacillans Larval ground colour has minimal contrast; subtly varying hues of dull green, subtly tinged purple in the subdorsal and dorsal regions; caudal horn thin, gently tapering in a shallow forward arc; (highly variable) .... ......................................... (Pl. 41) Macroglossum errans Larval ground colour has extreme contrast; subdorsal, dorsal and ventral regions are green, but the region between the subdorsal and subspiracular stripes is a combination of black, orange, and enlarged white dots; caudal horn stout, and a shallow ‘S’ shape (highly variable) ............................................................................... ...................... (Pl. 43) Macroglossum micacea micacea Key to pupae This key is specimen based and photographs may not be adequate for identification. It incorporates 70 species, those not included being unknown. It is preferable that specimens be examined under low magnification and preferably alive, although in most cases identifications can be obtained from cast skins. 1 – 2 – 3 – 4 – 5 – 6 – 7. – 8(1) Proboscis with a trunk-like extension that is free of the body (Figs 47, 49, 50) .............................................. 2 Proboscis without a trunk-like extension (Figs 48, 51, 52) ...................................................................................... 8 Trunk-like proboscis extension not recurved before its apex (Fig. 47) ................................................................... 3 Trunk-like proboscis extension recurved through some 180º or more (Fig. 49) .......................................... 6 In lateral view trunk-like proboscis extension reaching beyond level of dorsal distal margin of thorax .......... 4 In lateral view trunk-like proboscis extension shorter, not reaching to level of dorsal distal margin of thorax (Fig. 47) ........................................................................... 5 Proboscis with extension not projecting anterior of head; length including cremaster less than 55 mm ...... ............................................. (Pl. 53) Psilogramma argos Proboscis with extension projecting anterior of head; length including cremaster greater than 55 mm (2 species not distinguishable).............................................. (Pls 58, 57) Psilogramma papuensis or P. menephron nebulosa Abdominal segment 5 with a sharply deﬁned and prominent ridge immediately anterior of spiracle and which extends both dorsally and ventrally to spiracle . ............................................. (Pl. 37) Leucomonia bethia Abdominal segment 5 barely ridged immediately anterior of spiracle and only so dorsally to spiracle (3 species not distinguishable) ............................................. (Pls 54, 55, 56) Psilogramma casuarinae, P. exigua or P. maxmouldsi Proboscis extension recurved through more than 360º (Fig. 50) . ..................... (Pl. 14) Cerberonoton severina Proboscis extension recurved through some 180º (Fig. 49) ..................................................................................... 7 Total length of the proboscis extension if imagined as straightened out, reaching third abdominal segment (Fig. 49) .................................. (Pl. 7) Agrius convolvuli Total length of the proboscis extension if imagined as straightened out, reaching no further than second abdominal segment .................... (Pl. 8) Agrius godarti In dorsal view proboscis clearly forming the most anterior part of the head .............................................. 17 29/08/19 11:11:08.61 42 HAWKMOTHS OF AUSTRALIA Figs 47–64. Pupae. (47–52) lateral views. (53–64) terminal abdominal segments in dorsal view showing cremaster. – 9 – 10 In dorsal view proboscis not or barely extending anterior of head ............................................................... 9 Proboscis visible all the way to distal ends of wings .... .......................................................................................... 57 Proboscis not visible to distal ends of wings but partly hidden by legs, antennae and/or wing pads ............... 10 Small pupa, length clearly less than 35 mm, often much less. ...................................................................... 11 190613 Hawkmoths of Australia 3pp.indd 42 – 11 – 12 Larger pupa, length about 40 mm or more .............. 15 Proboscis not visible beyond distal ends of forelegs .... 13 Proboscis extending to or beyond distal ends of midlegs (hindlegs are not visible)................................ 12 Base of proboscis with a fused pair of rounded protuberances; non-glossy pupa (2 species not distinguishable) .................................................................. ............. (Pls 17, 18) Coenotes arida or C. eremophilae 29/08/19 11:11:09.27 THE AUSTRALIAN FAUNA – 13 – 14 – 15 – 16 – 17(8) – 18 – 19 – 20 – 21 – 22 – 23 – 24 – 25 – Base of proboscis without rounded protuberances; glossy pupa ................... (Pl. 59) Synoecha marmorata Abdominal segment 9 (the very narrow segment preceding last segment) not pitted or very weakly so .. .......................................................................................... 14 Abdominal segment 9 mostly pitted, similar to other abdominal segments ........ (Pl. 15) Chelacnema ochra Abdominal tergite 3 with pitting reaching spiracle ..... ...................................... (Pl. 34) Hopliocnema lacunosa Abdominal tergite 3 with pitting terminating well short of spiracle ...... (Pl. 34) Hopliocnema brachycera Very large pupa, 55 mm or more in length; dorsal surface of abdominal segments 1–7 entirely sculptured .......................................................................................... 16 Medium-sized pupa no longer than 50 mm in length; dorsal surface of abdominal segments 1–7 smooth and glossy except for anterior rim .......................................... ..................................................... (Pl. 36) Imber tropicus Distal part of proboscis covered by legs ......................... ....................................... (Pl. 20) Coequosa triangularis Proboscis visible to distal ends of legs ............................ ....................................... (Pl. 19) Coequosa australasiae Mesothorax with a distinct black or dark brown dorsal midline (totally black pupa should be treated as without such a line) ...................................................... 18 Mesothorax without a distinct black or dark brown dorsal midline ............................................................... 35 Keel-shaped proboscis long, in lateral view in front of head as long as or longer than its width (Fig. 51) .... 19 Keel-shaped proboscis short, in lateral view that part in front of head shorter than its width (Figs 48, 52) .... .......................................................................................... 20 Keel-shaped proboscis in lateral view with that part in front of head about as long as its width; cremaster in dorsal view longer than its maximum width................. ................................................. (Pl. 52) Nephele subvaria Keel-shaped proboscis in lateral view with that part in front of head clearly longer than its width; cremaster in dorsal view about as long as its maximum width .... ................................................... (Pl. 51) Nephele hespera Cremaster duck-bill shaped, apically broad in dorsal view (Figs 53–55, 58) ................................................... 21 Cremaster not duck-bill shaped, to the naked eye tending apically pointed in dorsal view (Figs 60–63).. .......................................................................................... 28 Duck-bill of cremaster without spines along lateral margin (view at x10) (Fig. 54). .................................... 23 Duck-bill of cremaster with one or more sharp lateral spines (Fig. 53). ............................................................. 22 Duck-bill strongly tapered in distal half (Fig. 53) (2 species not distinguishable) ........................ (Pls 44, 46) Macroglossum nubilum or M. prometheus lineata Duck-bill barely tapered in distal half (Fig. 55) ............ .............................. (Pl. 47) Macroglossum queenslandi Cremaster with much of dorsal surface smooth and glossy ............................................................................... 24 Cremaster with dorsal surface bearing multiple longitudinal grooves, or wrinkled ............................. 26 Small pupa with total length less than 40 mm ......... 25 Large pupa with total length greater than 40 mm ........ ............ (Pl. 39) Macroglossum corythus approximans Dorsal surface of abdominal segment 9 and cremaster both very glossy .... (Pl. 45) Macroglossum papuanum Dorsal surface of abdominal segment 9 far less glossy than cremaster .................................................................... .................................... (Pl. 50) Macroglossum vacillans 190613 Hawkmoths of Australia 3pp.indd 43 43 26 Proboscis anterior of head protruding about half the length of dorsal midline of head ................................ 27 – Proboscis anterior of head large, protruding more than length of dorsal midline of head, tending circular in lateral proﬁle ...... (Pl. 49) Macroglossum tenebrosa 27 Prothorax with many black dots (x10 magniﬁcation).. ...................... (Pl. 43) Macroglossum micacea micacea – Prothorax without black dots or just one or two .......... ...................................... (Pl. 42) Macroglossum joannisi 28(20) Cremaster with its pair of terminal spines projecting entirely laterally, i.e. sideways (Fig. 56) ..................... 29 – Cremaster with its pair of terminal spines directed distally in a V-shape or sometimes curved (Fig. 57) .... .......................................................................................... 31 29 Mesothorax with a black midline similar to that on abdominal segments ..................................................... 30 – Mesothorax with a black midline not similar to that on abdominal segments, only vaguely deﬁned in comparison ............. (Pl. 22) Daphnis placida placida 30 Abdominal segment 10 (the one supporting cremaster) laterally swollen near base; pupa usually about 70 mm in length, usually above 65 mm ....................................... ................................................... (Pl. 21) Daphnis moorei – Abdominal segment 10 (the one supporting cremaster) evenly tapering from base; pupa usually about 60 mm in length, usually below 65 mm ....................................... ...................... (Pl. 23) Daphnis protrudens protrudens 31 Cremaster in lateral view very short and very small; dorsally not glossy.............................................................. .......................... (Pl. 40) Macroglossum dohertyi doddi – Cremaster in lateral view long; glossy in dorsal view... .......................................................................................... 32 32 Abdominal segment 5 with its anterior ridge passing immediately anterior of spiracle; very small pupa, usually less than 35 mm in length ............................. 33 – Abdominal segment 5 with its anterior ridge passing through spiracle; larger pupa, usually more than 35 mm in length ................................................................. 34 33 Cremaster tapering to a pointed apex (Fig. 57). ........... ......................................... (Pl. 38) Macroglossum alcedo – Cremaster tapering to a broad apex (Fig. 58) ................ ................................. (Pl. 45) Macroglossum papuanum 34 Abdominal segment 7 with its dorsal posterior rim with pits boldly marked black; segment 8 with its anterior rim much smaller than posterior of segment 7; cremaster minutely pitted on apical quarter (x10 magniﬁcation) ............. (Pl. 48) Macroglossum rectans – Abdominal segment 7 with its dorsal posterior rim with pits not boldly marked black; segment 8 with its anterior rim about the same size as posterior of segment 7; cremaster smooth on its apical dorsal quarter (x10 magniﬁcation).............................................. ......................................... (Pl. 41) Macroglossum errans 35(17) Cremaster terminating in just a pair of simple spines (Figs 56, 64) ................................................................... 41 – Cremaster terminating in a pair of spines that are branched or more complex (Figs 59, 61) ................... 36 36 Cremaster with a pair of terminal spines that are bifurcate and have supplementary spines ................. 37 – Cremaster with a pair of terminal spines that are bifurcate but without supplementary spines ............ 38 37 Abdomen with subtle green patches; cremaster with hooked spines on and around base of the pair of the terminal branched spines and with many tubercles across the dorsal base of cremaster (Fig. 59) ................. ...................... (Pl. 24) Eupanacra splendens splendens 29/08/19 11:11:09.35 44 HAWKMOTHS OF AUSTRALIA – Abdomen entirely without green; cremaster with some large auxiliary spines on the pair of terminal branched spines but without tubercles across dorsal base of cremaster .................... (Pl. 33) Hippotion velox 38 Abdominal segment 10 (terminal segment) with dorsal surface ﬁnely granular, similar to cremaster ..... .......................................................................................... 39 – Abdominal segment 10 with dorsal surface closely pitted, contrasting with wrinkled cremaster ........... 40 39 Abdominal spiracles black but without a black surrounding highlight (x10 magniﬁcation). .................. ......................................... (Pl. 68) Theretra queenslandi – Abdominal spiracles accentuated by a surrounding jet black spot ..................... (Pl. 66) Theretra nessus nessus 40 Dorsal sculpturing on abdominal segment 8 (the segment bearing the caudal scar) much ﬁner than on dorsal surface of abdominal segment 10 (the last segment and the one bearing the cremaster) ................. ............................................. (Pl. 26) Gnathothlibus eras – Dorsal sculpturing on abdominal segment 8 similar to that on dorsal surface of abdominal segment 10 ...... ............................. (Pl. 25) Gnathothlibus australiensis 41(35) Cremaster terminating in a needle-like attenuation prior to apical spines (Fig. 56); not cavernous ventrally at base of cremaster ....................................................... 42 – Cremaster not terminating in a needle-like attenuation, clearly tapering evenly towards apex (Figs 62, 64); cavernous ventrally at base of cremaster .......................................................................................... 47 42 Proboscis with a distinct black blotch below eye and usually with caudal scar marked by a small black blotch............................................................................... 43 – Proboscis without a distinct black blotch below eye and without a black blotch on caudal scar ................ 46 43 Proboscis barely extending anterior of head in dorsal view (almost level with it), and not developed ventrally below head ............................. (Pl. 32) Hippotion scrofa – Proboscis clearly extending anterior of head and also developed ventrally below head .................................. 44 44 Dorsal surface of abdominal segment 1 ﬁnely and evenly pitted (x10 magniﬁcation) .................................... ........................................ (Pl. 27) Hippotion boerhaviae – Dorsal surface of abdominal segment 1 either without pitting or not evenly pitted, the pitting mainly conﬁned to the anterior margin ................................. 45 45 Length of pupa including cremaster 45 mm or longer .............................................. (Pl. 28) Hippotion brennus – Length of pupa including cremaster less than 45 mm (2 species not distinguishable) ......................................... ............... (Pls 28, 31) Hippotion brennus or H. rosetta 46 Many pits on dorsal abdominal surface highlighted black ........................................ (Pl. 32) Hippotion scrofa – Pits on dorsal abdominal surface not highlighted black ....................................... (Pl. 29) Hippotion celerio 47(41) Spines on cremaster with their bases close together, hence cremaster tends to have a narrow apex (x10 magniﬁcation) (Fig. 62) ............................................... 48 – Spines on cremaster with their bases wide apart, hence cremaster tends to have a broad apex (x10 magniﬁcation) (Fig. 64) .............................................. 54 48 Abdominal segments with dorsal and lateral surfaces ﬁnely granulated and mixed with ill-deﬁned pits (x10 magniﬁcation); never with a small, distinct, black spot subdorsally either side on most abdominal segments (not to be confused with lateral black markings around spiracles) ........................................................... 49 190613 Hawkmoths of Australia 3pp.indd 44 – 49 – 50 – 51 – 52 – 53 – 54 – 55 – 56 – 57(9) – 58 – 59 – 60 Abdominal segments with dorsal and lateral surfaces closely pitted with clearly deﬁned circular pits (x10 magniﬁcation); usually with a small black spot subdorsally either side on most abdominal segments.. ........................................ (Pl. 67) Theretra oldenlandiae Total length less than 55 mm, usually below 50 mm; dorsal surface of abdominal segment 8 closely and evenly pitted, distinct under magniﬁcation (x10).... 50 Total length usually 55 mm or longer, rarely below 55 mm; dorsal surface of abdominal segment 8 either not at all pitted or if so only at anterior and posterior margins ........................................................................... 53 A broad, dark, subdorsal band running almost length of body ............ (Pl. 69) Theretra silhetensis intersecta No subdorsal band ........................................................ 51 Ventral surface of abdominal segments 5 and 6 without a black midline.................................................... . .......................... (Pl. 40) Macroglossum dohertyi doddi Ventral surface of abdominal segments 5 and 6 with a black midline ................................................................. 52 Cremaster in dorsal view slender but not distally attenuated and needle-like (similar to Fig. 62) ............. ................................................ (Pl. 63) Theretra inornata Cremaster in dorsal view long, slender, needle-like (similar to Fig. 56) ............... (Pl. 29) Hippotion celerio Dorsal surface of abdominal segments with pitting along anterior margin of segments 3-7. .......................... ......................................... (Pl. 61) Theretra celata celata Dorsal surface of abdominal segments without pitting on any segments ................................................................. ....................... (Pl. 62) Theretra indistincta indistincta In dorsal view proboscis extending anterior of head about as far as head is long (along midline) ................... ..................................................... (Pl. 70) Theretra tryoni In dorsal view proboscis extending anterior of head much less than head is long ......................................... 55 Prothorax with its midline marked by a black line. ..... .............................. (Pl. 47) Macroglossum queenslandi Prothorax with its midline not marked by a black line .......................................................................................... 56 Pupa more than 45 mm long; abdomen gradually tapering to cremaster .......... (Pl. 64) Theretra latreillii Pupa less than 45 mm long; abdomen with an abrupt step-down in size between segments 7 and 8 ............... ................................................... (Pl. 71) Theretra turneri Head with a single, low rounded protuberance at base of proboscis that projects just enough to be the most anterior part of pupa .................................................... 58 Head without such a protuberance ............................ 61 Abdomen with a cavernous sublateral pocket each at junction of abdominal segments 2 and 3 and segments 3 and 4 ..................... (Pl. 4) Acosmeryx anceus anceus Abdomen without such cavities or with them barely developed, certainly not cavernous ........................... 59 Posterior margin of abdominal segment 7 wider than anterior margin of segment 8 so that there is a distinct step-down in size between the two segments ................ ...................................... (Pl. 5) Acosmeryx cinnamomea Abdominal segments 7 and 8 more or less conﬂuent apart from the usual depression between segments ..... .......................................................................................... 60 Maximum width of body rarely below 13.5 mm; in lateral view base of proboscis leaves head vertically .... ................................................ (Pl. 6) Acosmeryx miskini 29/08/19 11:11:09.45 THE AUSTRALIAN FAUNA – Maximum width of body rarely above 13.5 mm; in lateral view base of proboscis leaves head sloping backwards .................. (Pl. 5) Acosmeryx cinnamomea 61(57) Dorsal surface of abdominal segments evenly pitted or sculptured over entire surface (except intersegmental membrane) ................................................. 62 – Dorsal surface of abdominal segments coarsely pitted on about anterior quarter, much less so on remainder .......................................................................................... 72 62 Entire dorsal region of metathorax black (and often also much of pro- and mesothorax and usually also abdominal intersegmental membrane); otherwise light brown .............................. (Pl. 16) Cizara ardeniae – Entirely light to dark brown, sometimes almost entirely blackish ............................................................ 63 63 Glossy, very dark brown to blackish pupa (4 species not distinguishable) ........................................................... ...................................... (Pls 10–13) Cephonodes species – Not glossy or only slightly so, brown but never dark brown or blackish ......................................................... 64 64 Large pupae, length 40 mm or more ......................... 65 – Small pupae, length less than 40 mm ........................ 70 65 Cremaster terminating in a pair of spines that are branched apically .......................................................... 69 – Cremaster terminating in a pair of simple spines ....... 66 66 Cremaster in dorsal view long, slender, needle-like (Fig. 56) .................................. (Pl. 32) Hippotion scrofa – Cremaster not long, slender and needle-like (similar to Fig. 62) ....................................................................... 67 67 Mesothorax with a black midline similar to that on abdominal segments ..................................................... 68 – Mesothorax with a black midline not similar to that on abdominal segments, only vaguely deﬁned in comparison ............. (Pl. 22) Daphnis placida placida 68 Abdominal segment 10 (the one supporting cremaster) laterally swollen near base; pupa usually about 70 mm in length, usually above 65 mm ....................................... ................................................... (Pl. 21) Daphnis moorei – Abdominal segment 10 (the one supporting cremaster) evenly tapering from base; pupa usually about 60 mm in length, usually below 65 mm ....................................... ...................... (Pl. 23) Daphnis protrudens protrudens 69 Dorsal sculpturing on abdominal segment 8 (the segment bearing the caudal scar) much ﬁner than on segment 10 (the last segment and the one bearing the cremaster) .......................... (Pl. 26) Gnathothlibus eras – Dorsal sculpturing on abdominal segment 8 similar to that on segment 10......................................................... ............................. (Pl. 25) Gnathothlibus australiensis 70 Base of proboscis clearly anterior of eye crescent .... 71 – Base of proboscis about level with eye crescent............. ..................................................... (Pl. 72) Zacria vojtechi 71 Abdominal segments 5–7 with their anterior half or so raised into a low rounded ridge .................................. .............................................. (Pl. 35) Hyles livornicoides – Abdominal segments 5–7 similar in proﬁle to other abdominal segments ........ (Pl. 65) Theretra margarita 72(61) Anterior of head more or less rounded and conﬂuent . .......................................................................................... 73 – Anterior of head clearly divided into four rounded lobes....................... (Pl. 9) Angonyx papuana papuana 73 Small pupa less than 45 mm long; cremaster terminating in a long narrow pointed projection minutely bifurcate apically (x10 magniﬁcation) and without hair-like spines (4 species not distinguishable) ...................................... (Pls 10–13) Cephonodes species 190613 Hawkmoths of Australia 3pp.indd 45 – 74 – 45 Large pupa at least 45 mm long; cremaster bluntly rounded apically and much of its surface with minute hair-like spines .............................................................. 74 In lateral view antennae reach to ventral surface ......... ........................................... (Pl. 60) Tetrachroa edwardsi In lateral view antennae do not reach ventral surface (not ﬁgured) ................................... Acherontia lachesis Genus Acherontia [Laspeyres], 1809 Death’s head hawkmoths Type species: Sphinx atropos Linnaeus, 1758. By original designation. SYNONYMY Manduca Hübner, [1806]: [1]. Unavailable. A work rejected for nomenclatorial purposes by the International Commission on Zoological Nomenclature. Acherontia [Laspeyres], 1809: 100. Atropos Oken, 1815: 762. Unavailable. A work rejected for nomenclatorial purposes by the International Commission on Zoological Nomenclature. Brachyglossa Boisduval, 1828: 33. Atropos Agassiz 1846: 9. A junior homonym of Atropos Leach (a Psocoptera) and a junior objective synonym of Acherontia. Ochsenheimer (1808) divided the genus Sphinx Linnaeus into five parts which he called families. Later, in an anonymous 1809 review attributed by Ochsenheimer (1816) to Laspeyres, the four species included in Ochsenheimer’s ‘family IV’ were divided between two genera, three species to Sphinx, and Acherontia was proposed for atropos, thus establishing the genus. Worldwide: 3 species (Kitching and Cadiou 2000; Eitschberger 2003a; Kitching et al. 2018b). Australia: Acherontia lachesis (Fabricius, 1798). Mythology and superstition pervade the very essence of the almost universally recognised common name, death’s head hawkmoths. Kitching (2003) wrote ‘The death’s head hawkmoth, Acherontia atropos (Linnaeus, 1758), has the direst reputation of all hawkmoths, if not all Lepidoptera. This is due to the sinister-looking skull pattern on the thorax, and the Fig. 65. Acherontia lachesis, one of three species of Death’s head hawkmoths often associated with mythology and superstition. Photo Mark Hopkinson. 29/08/19 11:11:09.57 46 HAWKMOTHS OF AUSTRALIA Figs 66–67. Acherontia lachesis, Death’s head hawkmoth. (66) Last instar larva from Dauan Island, northern Torres Strait. Photo Cliff Meyer. (67) A Death’s head hawkmoth approaches a colony of Apis dorsata with the intent of robbing honey. The bees closest to the moth respond by shaking (dark pattern). Photo Nikolaus Koeniger. transverse black and yellow bands on the abdomen, which can be viewed as ‘ribs’. Add to this image the dark forewings, which at rest are draped on either side of the body like a cloak, and a high-pitched squeak emitted when the moth is disturbed, and the result is more than sufficient to inspire fear and dread in the uneducated and superstitious.’ The name Acherontia derives from Acheron, the River of Pain in the underworld of Greek mythology, and the sole species reaching Australia, A. lachesis, takes its name from the Fate who measures the length of the thread of life and determines destiny. Names of other Acherontia taxa are similarly conceived. The three known species have broad ranges, two of which overlap considerably (Kitching 2006). Acherontia atropos occurs through the Afrotropical region northwards as a migrant to the Palaearctic region and east to Iran. Acherontia lachesis (as two subspecies) and A. styx Westwood, 1847 (also as two subspecies) range through the Oriental region. Acherontia lachesis extends its range to New Guinea and northern Australia while A. styx reaches East Timor (Lane and Lane 2006). Adults are large, heavy and stout-bodied with a characteristic ‘death’s head’ or skull-like mark on the thorax. The proboscis is short, thick and ciliate, while the antennae are thick and straight with a terminal hook. The legs are short and thick with the mid and hindtarsi compressed and without a ventral bristle brush at the base. There is no pulvillus and the paronychium is reduced to a short lobe. The male genitalia have a long slender uncus but a very small, very short gnathos, the saccus terminates in a distally bulbous tubular extension, the juxta is broad and slightly tilted upwards either side of the midline; the valvae have a patch of long dentate friction scales, the harpes are apically bifurcate and usually claw-like and the phallus is long and very slender and without ornamentation. Both sexes can produce a squeaking sound that differs between species, with a more hiss-like sound in A. lachesis, a squeak-like sound in A. atropos and hoarser squeaks in A. styx (Kitching 2003). The sound is produced by air moved through the proboscis and is unique to Acherontia. It has twin origins, initially by the moth drawing air into the oral aperture at the base of the proboscis via a dilated pharynx which produces a rapid train of pulses, followed by expelling the air resulting in a brief sustained sound (Busnel and Dumortier 1959; Brehm et al. 2015). The process lasts only about 200 milliseconds and is repeated some 40–50 times to create the full audible squeak. Further, adults are able to detect high frequency sounds emitted 190613 Hawkmoths of Australia 3pp.indd 46 by bats through the palps in association with the pilifer (Göpfert and Wasserthal, 1999a, 1999b; Göpfert et al. 2002). Adult Acherontia are known for taking honey from bee hives (e.g. Tutt 1904; Newman 1965; Pittaway 1993; Koeniger et al. 2010) and their proboscis is modified for piercing rather than probing for nectar in flowers. But while honey may be a significant part of their diet, and attracts much attention, it is not their sole food source as they also feed on fermenting fruit (Choi et al. 2000; DAL pers. obs.) and nectar (Pittaway 1993). Based on circumstantial evidence, Kitching (2003, 2006) suggested that each of the three Acherontia species may be adapted to taking honey from specific Apis species. Indeed, all records of A. atropos taking honey from bee hives pertain to the European honey bee Apis mellifera. On the other hand, there are no records of A. styx or A. lachesis taking honey from Apis mellifera despite commercial farming of A. mellifera throughout the distributions of those two hawkmoths. There are multiple records of Acherontia lachesis taking honey from the giant honey bee Apis dorsata (Koeniger et al. 1999, 2010) and for Acherontia styx taking honey from Apis cerana (Koeniger et al. 2010). There is also a single record of Acherontia styx taking honey from Apis koschevnikovi (Koeniger et al. 2010) but Acherontia styx has never been record taking honey from Apis dorsata despite being common through the distribution of Acherontia styx. However, taking honey from bees is not without risk as bees vigorously defend their hive from intruders. Moritz et al. (1991) concluded that A. atropos renders itself ‘invisible’ upon entering the hive of Apis mellifera by chemically mimicking the bees’ cuticular fatty acids so that they smell like the bees, a highly specialized adaptation that suggests a long history of Acherontia-Apis interaction. Frances et al. (1985) found that Apis mellifera, A. cerana and A. dorsata each had different cuticular hydrocarbons. It follows then that A. styx and A. lachesis would need to mimic A. cerana and A. dorsata respectively if they are to rob those bees. Dreller and Kirchner (1995) indicated that honey bees have a well-developed sense of hearing. Rothschild (1985) suggested that the species-specific audible ‘squeaks’ produced by the three Acherontia species may also play a role in assisting the moths in avoiding detection while robbing hives but this has not been tested. Alternatively, Newman (1965) implied these sounds could be associated with mating, and Pittaway (1993) suggested they may be used to startle predators. Regardless, 29/08/19 11:11:09.71 THE AUSTRALIAN FAUNA included (subsequent combinations being omitted except for those relating to the name convolvuli), and the following unavailable names are omitted: alicea (Neuberger, 1899); extincta (Gehlen, 1928); fasciata (Pillich, 1909); fuscosignata Tutt, 1904; grisea Tutt, 1904; intermedia Tutt, 1904; major Tutt, 1904; minor Tutt, 1904; obscura Tutt, 1904; posticoconflua (Bryk, 1946); suffusa Tutt, 1904; unicolor Tutt, 1904; variegata Tutt, 1904; virgata Tutt, 1904. Sphinx convolvuli Linnaeus, 1758: 490 (type loc. not stated, [Europe]). Sphinx abadonna Fabricius, 1798: 435 (type loc. India Orientali). Herse convolvuli (Linnaeus): Oken, 1815: 762. Unavailable, rejected work (ICZN, 1956, 14: 3. Opinion 417). Agrius convolvuli (Linnaeus): Hübner, [1819]: 140. Sphinx patatas Ménétries, 1857: 90 (type loc. Taiti [=Tahiti]). Sphinx convolvuli roseafasciata Koch, 1865: 54 (type loc. New South Wales and Queensland). Sphinx pseudoconvolvuli Schaufuss, 1870: 15 (type loc. Port Natal [=Durban, South Africa]). Sphinx convolvuli var. distans Butler, 1874: 31, pl. 9 (type loc. New Zealand). Protoparce convolvuli (Linnaeus): Butler 1876b: 609. Protoparce distans (Butler): Butler, 1876b: 609. Protoparce orientalis Butler, 1876b: 609 (type loc. India, Scinde?, North Bengal, Moulmein, Ceylon, Hong-Kong, Java, Hakodadi). Macrosila convolvuli (Linnaeus): Behr, 1882: 3, 7. Sphinx convolvuli var. batatae Christ, 1882: 346 (type loc. not stated). Phlegethontius convolvuli (Linnaeus): Kirby, 1892: 690. Sphinx convolvuli var. nigricans Cannaviello, 1900: 295 (type loc. Eritrea). Herse convolvuli (Linnaeus): Rothschild and Jordan, 1903: 11, pls 22, 26, 35, 62, 63. Agrius convolvuli var. ichangensis Tutt, 1904: 333 (type loc. Ichang). Agrius convolvuli var. javanensis Tutt, 1904: 333 (type loc. Java). Agrius convolvuli var. tahitiensis Tutt, 1904: 333 (type loc. Tahiti). Protoparce convolvuli indica Skell, 1913: 56–61 (type loc. Sumatra). Herse convolvuli marshallensis Clark, 1922: 3 (type loc. Taluit, Marshall Islands). Herse convolvuli peitaihoensis Clark, 1922: 2–3 (type loc. Pei-tai-ho [=Beidaihe], northern China) Herse convolvuli aksuensis O. Bang-Hass, 1927: 78 (type loc. Tian Shan, China) Rougerie et al. (2014) in their study of DNA barcodes found that Agrius convolvuli was divided into two distinct genetic groupings, one widespread west of the islands of South-East Asia in Europe, Africa, India to China, the other one widespread east of the islands of SouthEast Asia, east of the Palau Islands and Moluccas, through New Guinea, Australia, New Zealand and eastern Pacific islands. The COI barcodes of these two groups showed a significant divergence of 3.32%. The western group has an internal maximum variance of 1.73%, and the eastern group’s internal maximum variance is about 1.55%. Although this strongly suggests speciation, the two clusters are separated by a large geographic gap from where few samples have been reported. There are no sequenced A. convolvuli samples from mainland South-East Asia, the Malaysian archipelago and only a single specimen from Indonesia. Consequently, it is not yet possible to determine if the two groups retain their separate identities across this geographically intermediate region. Except for size, with western specimens being on average a little larger than the eastern, we could find no consistent differences in colour, wing markings, or morphology (including proboscis length, and male and female genitalia) between the two gene types. Given the uncertainty about the perceived genetic difference between the two A. convolvuli gene types and because we could find no other discernible difference in the morphology of adults or the early stages (c.f. images in Eitschberger and Ihle 2014: 192, 199, 200), we refrain from distinguishing the two genetic groups taxonomically in the absence of additional evidence. DISTRIBUTION AND HABITAT Cocos-Keeling Islands (Holloway [1983]; Lachlan 2006b) and Christmas Island (Moulds 1986) in the Indian Ocean, widespread throughout mainland Australia and in Tasmania. It is also found on most offshore islands, including Barrow 190613 Hawkmoths of Australia 3pp.indd 59 59 Island in Western Australia, Melville and Bathurst Islands (the Tiwi Islands) in the Northern Territory, Dauan, Hammond and Yam Islands in Torres Strait, Lizard, Dunk and remote Willis Island in the Coral Sea (Farrow 1984), Lord Howe and Norfolk Islands (Holloway 1977) in the Tasman Sea and King Island in Bass Strait (Martyn et al. 1972). The species inhabits a wide range of habitats from dry inland areas to the wet tropics. Savannah woodlands and subtropical coastal regions are often favoured wherever Convolvulaceae are growing. Beyond Australia, A. convolvuli is one of the most broadly distributed sphingids in the world with a range that extends throughout Europe, Africa, Asia, and the islands of the Pacific Ocean as far east as the Marquesas and Pitcairn Islands. ADULT DIAGNOSIS Sexually dimorphic. Proboscis 78–108 mm long. Abdomen with a lateral row of distinct pinkish red patches. Forewings 32–49 mm long (male), 33–51 mm (female), the females usually larger than males and specimens from arid regions sometimes unusually small. Male forewings above grey strongly patterned with complex black markings variable between individuals but usually incorporating a large black patch on costal margin about midlength; discal spot grey, sometimes distinct, sometimes not discernible. Female forewings above grey with only subtle, very fine, black markings. Both sexes have the hindwings above with a black subbasal band or blotch, two medial diffused black bands sometimes merging, and a similar one subterminal, all variable between individuals. Wings below grey to light brown with few markings. There are small variations in wing and body colour that have led to the naming of many forms and varieties, especially in Europe. Male genitalia (Pl. 84, fig. e) with uncus apically sharply pointed and laterally dilated subapically; gnathos long and narrow, almost as long as uncus, very lightly sclerotised, almost flat, very thin and very flexible so that it follows the anal tube on its inversion during dissection; saccus short, gradually tapering to a broadly rounded to truncate apex; juxta cylindrical, shorter than wide, lower half strongly sclerotised with broad, upturned, lateral wings that meet a lightly sclerotised membranous upper half; valvae with a patch of small short spines beyond harpes; harpes short, broad, flat, apically with an upcurved club-shaped appendage and a longer adjacent upcurved subapical spine-like appendage; phallus short, without ornamentation, apically bevelled from ventral surface. 29/08/19 11:11:11.15 60 HAWKMOTHS OF AUSTRALIA Agrius convolvuli is similar to A. godarti from which it can be distinguished by its abdominal markings pinkish red rather than buff yellow, and by the four black hindwing bands with the middle two sometimes merged in contrast to the three distinct bands of A. godarti. The male genitalia clearly differ in lacking a robust conical projection on the ventral margin of the valvae just beyond the harpe. DESCRIPTIONS OF IMMATURE STAGES Egg (Pl. 7, fig. a) Pale bluish green darkening to light brownish yellow prior to hatching; glossy; at x50, closely covered with shallow circular depressions; ovoid; unusually small for a hawkmoth of its size, 1.3–1.4 mm long x 1.1–1.2 mm wide x 1.0–1.2 mm high. Duration usually 3–4 days. Larva (Pl. 7, figs b–j) Usually there are five instars but under stressed conditions A. convolvuli may proceed to a sixth instar. Eitschberger and Ihle (2014) illustrate larvae from Thailand. First instar (Pl. 7, figs b, c). Pale yellowish on hatching becoming pale green darkening as feeding progresses but usually remaining pale green or pale yellow on thorax and distal abdominal segments; semi-glossy; primary setae very fine and not easily discernible (x25), microscopically bifurcate at extreme apex, some setae on anal plate longest and arising from conical tubercles about as high as wide; prothoracic shield tending pale yellow, inconspicuous; spiracles tending pale brown, inconspicuous. Head pale yellow to pale green, without markings. True legs pale yellow to very pale grey; ventral prolegs, claspers and anal plate pale yellow to pale green. Caudal horn black, sometimes with traces of brown at base; slender and long (1.5–1.6 mm); gently conical; straight or nearly so; surface of horn and apical branches with numerous spine-like black tubercles (x25); apex broadly bifurcate, the branches short, each with a long, fine, seta. Length on hatching 3.7–4.0 mm; length at maturity 8–9 mm. Width of head capsule about 0.7 mm. Duration usually 3–4 days. Second instar (Pl. 7, fig. d). Some individuals a little glossy, others without gloss; light green with dorsal midline distinctly darker green; a suggestion of a pale subdorsal band from head to base of caudal horn; dorsal and lateral surfaces of thorax and abdominal segments 1–7 with numerous small tubercles in transverse rows each with a fine pale seta minutely bifurcate at apex (x100); abdominal segment 8 with similar tubercles and setae but not all arranged in regular transverse rows; spiracles black or tending so (rarely pale), those on abdominal segments 7 and 8 considerably larger than remainder and elliptical in shape, the smaller spiracles circular. Head without markings; green with many tubercles and setae similar to those of body. True legs pale brown, glossy; ventral prolegs from entirely light green to very pale brown with grey to black lateral shields; anal plate green with tubercles and setae similar to those of body. Caudal horn black, usually dull pinkish at base on lateral surfaces (rarely with base entirely dull pinkish); straight, gently conical, slender and long (2.2–3.0 mm); densely covered by short, conical tubercles coloured similar to adjacent part of the horn, each with a fine seta; apex of horn bifurcate, each branch short, conical, and terminating in a fine seta, the branches spreading about as wide as distal part of horn. Length at maturity 14–16 mm. Width of head capsule 1.1– 1.2 mm. Duration usually 3–5 days. 190613 Hawkmoths of Australia 3pp.indd 60 Third instar (Pl. 7, fig. e). Light yellowish green to light bluish green; with or without black markings; dorsal midline narrowly dark green, indistinct on thorax; an indistinct subdorsal pale yellow stripe clearest on thorax; those larvae lacking or with minimal black markings with seven thin, pale yellow, oblique lateral stripes on abdominal segments 1–8, these often indistinct (rarely absent), the lower end of these stripes vague, below spiracular line and more or less reaching posterior margin of preceding segment, each stripe inclined backwards from base in a straight line but usually gently curved before terminating near dorsal midline, the most posterior stripe terminating at base of caudal horn; spiracles always black, often narrowly circled black. Head without markings; pale green. True legs light brown with black lateral shields; ventral prolegs, claspers and anal plate similar in colour to adjacent body, sometimes with black patches on prolegs. Dark form similar to 4th instar but black never intense. Caudal horn either black as in 2nd instar or multicoloured as in 4th instar; barely curved forwards, slender, gradually tapering to a pointed apex; 3.5–5.0 mm long; densely covered by short, conical tubercles coloured similar to adjacent part of the horn, each with a fine short seta. Length at maturity 24–26 mm. Width of head capsule 1.8–1.9 mm. Duration usually 3–6 days. Fourth instar (Pl. 7, figs f, g). In both green and dark forms. Green form light yellowish green to light bluish green; with or without black markings; dorsal midline vaguely darkened; thorax with an indistinct subdorsal pale yellow stripe sometimes extending a little onto abdomen; those larvae without black markings with seven thin, pale yellow, oblique lateral stripes on abdominal segments 1–8, usually indistinct (rarely absent), their lower end below spiracular line and more or less reaching posterior margin of preceding segment, each stripe inclined backwards from base in a straight line but usually gently curved before terminating near dorsal midline, the most posterior stripe terminating at base of caudal horn; spiracles orange, partially or entirely circled black. Head with a pale yellowish stripe on each cheek from vertex to base of antenna, also sometimes with a prominent black stripe adjoining pale yellowish stripe along its outer margin. True legs blackish, glossy. Dark form either entirely black or partly black, the latter mostly dorsal and never obliterating the yellow, oblique lateral stripes; body with a subdorsal yellow band clearly separating the dorsal and lateral areas and a similar but narrower yellow band sublaterally. Caudal horn either black often with reddish brown and sometimes pale yellow at base, otherwise multicoloured with the apical tip black adjoined by bright yellow for about another quarter with the basal half or so pale green to greyish green, the junction between the yellow and green merging and indistinct, the dorsal midline black around midlength and brownish on basal quarter or so; straight or barely curved forwards in a sweeping arc; gently tapering to a pointed apex sometimes minutely bifurcate, long (5.0–7.0 mm); densely covered by short, spine-like distally directed tubercles just visible to naked eye, similar in colour to adjacent part of horn. Length at maturity 38–41 mm. Width of head capsule 2.8–3.0 mm. Duration usually 4–7 days. Fifth instar (Pl. 7, figs h–j). Colour and markings show considerable variability between individuals but basically there are two colour forms, a green form in various shades and with or without black markings, and a dark form in shades of brown or black; not glossy. Green form sometimes with an irregular black subdorsal stripe from head to base of caudal horn, the anterior ends converging; abdomen with seven black, oblique, lateral stripes with their lower margins sometimes 29/08/19 11:11:11.21 THE AUSTRALIAN FAUNA edged white, pale yellow or greenish, the lower end of each stripe anterior of a spiracle and thereafter inclined backwards to dorsal surface before fading, the most posterior stripe terminating at base of caudal horn; spiracles white and black to varying degrees, but usually highlighted by a black blotch and with a white circumference, sometimes entirely black. Spiracles black but in some individuals with lateral margins white. Prothoracic shield indistinct, lightly sclerotised, without noticeable tubercles even at x25, semi-glossy, pale green sometimes with black around midline. Head semiglossy; green, similar to body, always a black stripe down each cheek from vertex to base of antenna, sometimes a similar second stripe down each cheek more medially and less intensely black or pale yellow, sometimes a narrow black stripe following the centre of head forking along adfrontal sutures; antennae mostly pale green; mouthparts mostly black. True legs glossy black; ventral prolegs blackish with lateral shields glossy black and lower margin brown; claspers green with variable black markings; anal plate green with white or pale yellow edging and sometimes variable black markings; semiglossy. Dark form dull black or various shades of brown; a white or pale yellow, broken, subdorsal stripe from head to base of caudal horn; a whitish irregular, sublateral stripe from head to anal plate, usually more prominent than subdorsal stripe; seven white or pale yellow, narrow, oblique, lateral stripes, one each on abdominal segments 1–7, each diagonally across the width of the segment between the subdorsal and sublateral stripes and extending, to some degree, through the subdorsal stripe across the intersegmental membrane, in black larvae these oblique stripes fragmented and sometimes almost absent; body ventrally similar to dorsal and lateral surfaces. Spiracles black but in some individuals with lateral margins white. Prothoracic shield semi-glossy, brown or black, but often with pale yellow or pale orange either side in a continuation of the subdorsal stripes; with scattered, minute tubercles visible under magnification, barely rising above surface. Head semi-glossy; light brown to brownish orange; two black stripes down each cheek from vertex to antenna, the outer stripe broadest and tapering towards vertex; a somewhat similar black stripe at rear of each cheek usually obscured by prothorax; a narrow black stripe down centre of head forking along adfrontal sutures; antennae mostly pale yellow; mouthparts mostly black. True legs black, tending glossy; ventral prolegs blackish with lateral shields glossy black with lower margin brown; claspers black; anal plate black with white, pale yellow or pale brown edging, semi-glossy. Caudal horn of both green and dark larvae semi-glossy; usually orange or reddish brown at least with apical quarter distinctly black but on some black larvae horn entirely black; curved backwards in an arc to equal about a quarter circle or a little less; tapering evenly to a pointed apex; 6–9 mm long; with many scattered, spine-like tubercles directed distally, very short, each with a short, simple seta, these tubercles similar in colour to adjacent part of horn. Length at maturity 85–105 mm. Width of head capsule 4.5–5.2 mm. Duration usually 8–9 days. Pupa (Pl. 7, figs k–m) Semi-glossy; varying shades of brown but usually tending reddish brown, usually a little paler ventrally, without distinct markings although often with some darkening primarily on head, metathorax and around anterior and posterior regions of abdomen. Proboscis with a trunk-like extension that has its distal third or so sharply recurved through 180º or more to adjoin or almost meet the ventral surface of thorax, coloured brown to nearly black; proboscis along ventral midline brown 190613 Hawkmoths of Australia 3pp.indd 61 61 and usually tipped black. Head, prothorax and mesothorax at x10 finely rugose. Wings glossy, smooth, with veins barely raised; brown, sometimes with a faint greenish tinge weakening with maturity, sometimes edged black along distal margin. Antennae and legs brown, often becoming blackish with maturity. Abdominal segments 1–8 finely pitted, especially on anterior quarter, smoother on their posterior quarter or finely rugose; abdominal segment 9 entirely rugose on dorsal and lateral surfaces, less so ventrally; spiracles distinct, black but lacking a black surround; spiracular furrows with first ridge anterior of spiracle strongly developed, sometimes with secondary ridges before cavity. Cremaster black, sometimes reddish brown basally; in dorsal view more or less an equilateral triangular, coarsely granulated, with adjoining abdominal surface folded longitudinally; in lateral view tapering to a point along dorsal margin; ventral surface similar in texture to dorsal surface, depressed along midline; apex in dorsal view terminating in a pair of short, sharp spines in a V-shape, their slender apices very fine and usually missing in handled specimens. Length 50–55 mm. Width at widest point 11.5–12.5 mm. Duration usually 2–3 weeks during warmer months while late season pupae overwinter. BIOLOGY Larval foodplants CONVOLVULACEAE: Calystegia sepium (Jenny Holmes); *Ipomoea aquatica kang kong, *I. batatas sweet potato, *I. cairica, *I. hederifolia, *I. indica morning glory, *I. nil; *Merremia dissecta; Polymeria sp. (Byrne and Byrne 2008); Stictocardia tiliifolia (TDS). It is a minor pest of sweet potatoes in New Guinea where this is a food staple (Szent-Ivany 1958). In captivity, larvae will accept almost any Convolvulus species. Zagorinskii et al. (2013) successfully reared larvae on an artificial diet. In an area north-west of Atherton, northern Queensland, eggs have been found on three occasions deposited on the undersides of juvenile leaves of Clerodendrum floribundum (Lamiaceae) in open eucalypt woodland. While all the eggs hatched, the larvae refused to eat Clerodendrum and perished. Moulds (1981) lists Abutilon oxycarpum (Malvaceae), but this record is now considered erroneous and was probably based on a wandering larva. Egg Laid singly on the foodplant, either on upper or underside of juvenile leaves and occasionally on stems. Larva Larvae are docile and not overly responsive to disturbances. Throughout much of the range of A. convolvuli, larvae are mostly found during November to May rains although in some areas, larvae can be found throughout the year. Martyn et al. (1973, 1977) record larvae in Tasmania during February and May. Early instar larvae rest on the undersides of juvenile leaves of the foodplant and feed from the leaf margin. Late instar larvae tend to rest on stems or under large leaves within lower sections of their foodplant vines and are remarkably well camouflaged. At Darwin during March, larvae of all instars were common in a large patch of Ipomoea nil, which was regularly inspected to observe larval behaviour. Among nearly two hundred larvae in an area estimated at 100 m2, both green and brown 4th and 5th instar larvae appeared in approximately equal numbers. The green larvae fed both during the day and 29/08/19 11:11:11.28 62 HAWKMOTHS OF AUSTRALIA night, but brown or black larvae only fed during the night and remained hidden during the day in lower sections of their foodplant or in the shrubs supporting the vines. At that time, maximum daytime temperatures were high (35–38ºC), with minimum temperatures of 27–28ºC overnight. Bell and Scott (1937) and Pittaway (1993) comment on similar behaviour in larvae from India and the western Palaearctic. Pupation Prepupal larvae become very mobile and can wander 100 m or more in search of a suitable location to pupate. Pupation occurs in an underground cell of well-packed earth up to 15 cm below ground level, without silk on the walls. The pupa is capable of vigorous side-to-side or circular abdominal movement. a large movement of adults in September 1982. After only seeing one adult for several weeks, 40 individuals came to the light in a single night, 14 came the next night and subsequently just a few singletons. In the eight years of monitoring his light every night, the incident was never repeated. In warmer years, individuals move south to Tasmania but populations that establish there eventually succumb to harsh winters. It is also an intermittent migrant to New Zealand and sometimes breeds there (Hudson 1898, 1928; Gaskin 1970; Dugdale 1988). Fox (1978) showed that migration of Australian Lepidoptera to New Zealand was primarily dependent upon lows developing in the southern Tasman Sea causing strong and prolonged westerly winds or from tropical cyclones causing similar winds when they move sufficiently south. Parasitoids and predators TACHINIDAE: Blepharipa sp. (Queensland Museum); Sturmia convergens (from Crosskey 1973). Adult Throughout Australia, adults are mostly encountered during the warmer months of the year from late spring to early autumn although in the tropical north they can be found in all months. In the arid interior, adult emergence follows heavy rains in association with hot weather, in some years in large numbers. It is a common species throughout most of its distribution and is attracted to light throughout the night, but especially after midnight. There are multiple generations under favourable conditions. Harbich (1980b) found that several generations could be hand reared in a year. Adults feed from tubular flowers at dusk and early evening. In the Cairns region in northern Queensland, lantana, spider lilies (Hymenocallis sp.) and male pawpaw (Carica papaya) are favourites. In Toowoomba, lantana, jacaranda and abelia are frequented. At times feeding adults can be numerous and on one occasion near Torrens Creek west of Charters Towers, adults were in such numbers that they could be heard hovering as they fed from eucalypt blossom at dusk. Adults have defined migratory patterns in the northern hemisphere summer in Africa, India, South-East Asia and southern Europe, and from southern Europe regularly extend those flights into Scandinavia and sometimes beyond into far eastern Russia. Migratory patterns in Australia are little understood but there is no evidence of migration north out of Australia into the wet tropics of Indonesia or New Guinea. The wet tropics of Malaysia, Indonesia and New Guinea may be a barrier to individuals from the northern hemisphere moving south and to those in the southern hemisphere moving north. This may be because rainforests provide few larval foodplants for breeding and migration seems to be, at least in part, associated with expanding into cooler regions as they warm during summer so that northern hemisphere adults migrate north while southern hemisphere adults move south. There are two observations that strongly suggest A. convolvuli migrates in Australia. At Atherton, northern Queensland, large numbers of adults were seen at streetlights on two occasions four years apart during August, each time over just three nights, with numbers sharply declining over the second and third nights, with none seen thereafter. Although direction of flight could not be determined, during the same period, no adults were seen at Mareeba, some 35 km north. The second account comes from an extensive light trapping study over eight years by Peter Mackey (pers. comm.) in Rockhampton, Queensland, when he encountered 190613 Hawkmoths of Australia 3pp.indd 62 Agrius godarti (W.S. Macleay, 1826) Adults, Pl. 8, fig. l; Pl. 73; Pl. 84, fig. f. Immatures, Pl. 8, figs a–k. SYNONYMY Sphinx godarti W.S. Macleay, 1826: 464 (type loc. not stated, northern Australia). For a detailed explanation relating to this type locality and the type specimen see under synonymy of Cephonodes kingii. Diludia? godarti (W.S. Macleay): Butler, 1876b: 615. Sphinx distincta Lucas, 1891a: 4 in original issue; 894 when republished in The Queenslander (type loc. near Kimberley, North Australia). Phlegethontius? distincta (Lucas): Kirby 1894: 102. Herse godarti (W.S. Macleay): Rothschild and Jordan, 1903: 9, pl. 35. Agrius godarti (W.S. Macleay): D’Abrera, [1987]: 12. DISTRIBUTION AND HABITAT Australian endemic found throughout most of continental Australia, but as yet unknown from the south-west of Western Australia. It is essentially a dry country species preferring semi-arid and arid habitats although at times it is also found in wetter areas including tropical north-eastern Queensland. Southern records from Western Australia include Carnarvon, Kalgoorlie and the Madura district. In Queensland, there are no records north of Laura except for a single old specimen in the SAM labelled as coming from Moa Is, a doubtful locality requiring confirmation. From western New South Wales, there are no records south of Broken Hill but along the coast A. godarti ranges at least as far south as Wallaga Lake, where it is regularly encountered (Glenn Cocking pers. comm.), although it is only an occasional visitor to Sydney. There is a single specimen in the ANIC from Canberra, A.C.T. The species occurs in north-western Victoria with records from Mildura, Rainbow and Lake Hindmarsh (Fabian Douglas pers. comm.), and there is an old record from Coburg, a northern Melbourne suburb. There are few records from South Australia although the species appears to be widespread and occurs south to Adelaide. No specimens are known from Tasmania. The primary habitat is dry savannah woodland. Yet, A. godarti regularly appears in wet areas of north-eastern and south-eastern Queensland, such as the Atherton Tablelands and Lamington Plateau, although it does not appear to establish breeding populations in those areas. ADULT DIAGNOSIS Male and female similar. Proboscis 62–76 mm long. Abdomen with a lateral row of distinct yellowish patches. Forewings 34– 46 mm long (male), 36–49 mm (female), the females usually 29/08/19 11:11:11.34 THE AUSTRALIAN FAUNA larger than males and specimens from arid regions are sometimes unusually small; above grey strongly patterned with complex blackish markings variable between individuals but usually incorporating a large blackish patch on costal margin about midlength; discal spot whitish, always visible. Hindwings above with a black subbasal band, one medial and a similar one subterminal, all well defined but a little variable between individuals. Wings below grey to light brown with few markings. Male genitalia (Pl. 84, fig. f) with uncus apically sharply pointed and laterally dilated subapically; gnathos long and narrow, almost as long as uncus, very lightly sclerotised, almost flat, very thin and very flexible so that it follows the anal tube on its inversion during dissection; saccus short, gradually tapering to a broadly rounded to truncate apex; juxta cylindrical, shorter than wide, on its lower half strongly sclerotised with broad, upturned, lateral wings that meet a lightly sclerotised membranous upper half; valvae with a patch of small short spines beyond harpes that reach a robust conical projection on ventral margin beyond harpes; harpes short, broad, flat, with two large parallel upcurved sharp hook-like appendages; phallus short, without ornamentation, apically bevelled from ventral surface. Agrius godarti is similar to A. convolvuli from which it can be distinguished by its lateral abdominal markings, which are yellowish rather than pinkish red, and by its well defined three black hindwing bands rather than four with the middle two sometimes merged. The male genitalia clearly differ in having a robust conical projection on the ventral margin of the valvae beyond the harpe. DESCRIPTIONS OF IMMATURE STAGES Egg (Pl. 8, fig. a) Varying shades of green becoming dull just prior to hatching; glossy; at x25 with a fine reticulate pattern of shallow circular depressions; ovoid; unusually small for a hawkmoth of its size, 1.1–1.2 mm long x 1.1–1.2 mm wide x 0.9–1.0 mm high. Duration usually 3–4 days. Larva (Pl. 8, figs b–h) First instar (Pl. 8, figs b, c). Pale yellowish on hatching, becoming pale greenish, sometimes a little paler on anterior thorax and distal abdomen; semi-glossy; primary setae very fine and very short and not easily discernible (x25), microscopically bifurcate at extreme apex, most arising from minute black tubercles, some setae on anal plate longest and 190613 Hawkmoths of Australia 3pp.indd 63 63 arising from conical tubercles about as high as wide; prothoracic shield tending pale yellow, inconspicuous; spiracles tending pale brown, inconspicuous. Head pale green to pale yellow, without markings. True legs pale, usually brownish; ventral prolegs, claspers and anal plate similar in colour to body; anal plate with a median pair of slightly larger tubercles. Caudal horn black, sometimes with traces of brown at base; straight or nearly so; almost parallel-sided, barely tapering to a blunt point, 1.5–1.7 mm long; numerous microscopic tubercles barely discernible at x50; apex broadly bifurcate, the branches short, each with a long, fine seta. Length on hatching about 3.5 mm; length at maturity about 7–8 mm. Width of head capsule about 0.7 mm. Duration about 3 days. Second instar (Pl. 8, fig. d). Sometimes a little glossy; light green with dorsal midline darker green; a pale yellow or whitish subdorsal band from head to base of caudal horn, sometimes indistinct; dorsal and lateral surfaces of body with numerous, very small pale tubercles in transverse rows each with a fine pale seta minutely bifurcate at apex (x100); spiracles brown or black, sometimes surrounded by a small black blotch, spiracles on abdominal segments 7 and 8 considerably larger and elliptical rather than circular. Head without markings; green to pale brown with many tubercles and setae similar to those of body. True legs pale yellow to pale brown, glossy; ventral prolegs and claspers light green to very pale brown; anal plate pale green with a median pair of slightly larger tubercles. Caudal horn pale pink to dark red to mostly black, often a mixture of these colours; straight, gently conical, slender, 1.9– 2.0 mm long; densely covered by short, conical tubercles coloured similar to adjacent part of the horn, each with a fine seta; apex minutely bifurcate, the branch short, conical, terminating in a fine seta, and spreading about as wide as distal part of horn. Length at maturity 16–18 mm. Width of head capsule 1.1– 1.3 mm. Duration usually 3–4 days. Third instar (Pl. 8, fig. e). Light green, with or without black markings variable between individuals; a pale yellow subdorsal stripe from head to base of caudal horn usually distinct; abdominal segments 1–8 usually with seven thin, pale yellow, oblique lateral stripes, the lower end of each often vague, sometimes reaching posterior margin of preceding segment, each stripe inclined backwards to terminate at or a little beyond the pale subdorsal stripe, the most posterior stripe terminating at base of caudal horn; at x25 dorsal and lateral surfaces of body with numerous, very small pale tubercles in transverse rows; spiracles brown or black, rarely orange, often narrowly circled black; prothoracic shield indistinct, similar in appearance to remainder of thorax. Head green, often with a pale stripe down each cheek. True legs pale pink to light brown, sometimes tending black; ventral prolegs, claspers and anal plate similar in colour to adjacent body, the anal plate with a pale margin. Caudal horn pale pink to dark red to mostly black, sometimes with a little pale yellow, often a mixture of these colours; straight or barely curved forwards; slender, about 3 mm long; gradually tapering to a pointed apex; densely covered by short, conical tubercles coloured similar to adjacent part of the horn, each with a fine short seta. Length at maturity 28–29 mm. Width of head capsule 1.8–2.0 mm. Duration about 3 days. Fourth instar (Pl. 8, fig. f). In both green and dark forms. Green form light yellowish green to light bluish green; with or without black markings; not glossy; a white or pale yellow subdorsal stripe from head to base of caudal horn, often broken between segments, sometimes with black markings 29/08/19 11:11:11.50 64 HAWKMOTHS OF AUSTRALIA along upper margin; dorsal midline marked by a very thin dark bluish green line; abdominal segments 1–8 with seven thin, white or pale yellow, oblique lateral stripes, these sometimes indistinct, the lower end of each level with or below spiracular line and more or less reaching posterior margin of preceding segment, each stripe inclined backwards from base in a straight line to merge with a similarly coloured subdorsal stripe, the most posterior stripe terminating at base of caudal horn; many transverse bands of minute pale tubercles visible under magnification, these tubercles short, conical and each with a very minute short simple seta; spiracles orange, often partially or entirely circled black; prothoracic shield indistinct, coloured similarly to rest of thorax. Head green with a pale yellow or white stripe down each cheek from vertex to base of antenna, also sometimes with a prominent black stripe adjoining pale yellowish stripe along its outer margin; with many scattered tubercles similar to those of body; mouthparts green and brown or mostly black. Legs glossy, light dull green to brown or sometimes black; ventral prolegs, claspers and anal plate similar in colour to body, the anal plate usually with a pale margin. Dark form with markings similar to green form, the body black to varying degrees. Caudal horn of both colour forms black often with brown and sometimes pale yellow at base; straight or nearly so; 5.5– 6.0 mm long; gently tapering to a pointed apex sometimes minutely bifurcate; densely covered by short, distally directed, spine-like tubercles just visible to naked eye similar in colour to adjacent part of horn. Length at maturity 40–41 mm. Width of head capsule 3.2–3.3 mm. Duration usually 4–8 days. Fifth instar (Pl. 8, figs g, h). The colour and markings between individuals show considerable variation but basically there are two colour forms, a green form in varying shades and with or without black markings, and a dark form in shades of brown or black; not glossy. Green form light greenish yellow to brownish yellow to green, usually with bold black markings; a white or pale yellow subdorsal stripe from head to base of caudal horn but often overlaid in part or completely by black that is either intermittent or continuous; a similar white or pale yellow sublateral stripe also sometimes partially or completely overlaid by black; abdomen with seven, white or pale yellow, oblique, lateral stripes, also often partly or entirely overlaid by black, the lower end of each stripe anterior of a spiracle and thereafter inclined backwards to dorsal surface before fading, the most posterior stripe terminating at base of caudal horn. Spiracles orange, usually surrounded by black. Prothoracic shield indistinct, lightly sclerotised with many minute tubercles and tending semiglossy, coloured similarly to adjacent body. Head green with a pale yellow or white stripe down each cheek from vertex to base of antenna, also usually with a prominent black stripe adjoining pale yellowish stripe along its outer margin; mouthparts green and brown or mostly black. True legs glossy, black or brown and black; ventral prolegs, claspers and anal plate green, brown or sometimes blackish, often a mixture of each but anal plate usually with a pale margin and the claspers and anal plate mostly semi-glossy. Dark form black or various shades of brown; a whitish or pale yellow subdorsal stripe from posterior margin of prothoracic shield to base of caudal horn, on most larvae extensively interrupted by faded and missing sections; a pale yellow or very pale green irregular sublateral stripe from head to anal claspers, usually more prominent than subdorsal stripe; seven whitish or pale yellow, narrow, oblique, lateral stripes, one each on abdominal segments 1–7, each diagonally across the width of 190613 Hawkmoths of Australia 3pp.indd 64 the segment between the subdorsal and sublateral stripes, but also extending to some degree through the subdorsal stripe across intersegmental membrane, these oblique stripes often fragmented and sometimes almost absent; ventral surface of body tending a little paler than remainder. Spiracles orange and sometimes ringed black. Prothoracic shield semi-glossy, brownish orange to black but often with pale yellow or pale orange either side in a continuation of the subdorsal stripes; with scattered, minute tubercles visible under magnification, these barely rising above surface. Head semi-glossy; light brown to brownish orange; two black stripes down each cheek from vertex, or just below vertex, their bases one on each side of antenna, these stripes tapering towards their extremities; a similar black stripe at rear of each cheek usually obscured by prothorax; a narrow black stripe down centre of head forking along adfrontal sutures; antennae mostly pale yellow; mouthparts mostly black. True legs glossy black; ventral prolegs blackish with lateral shields glossy reddish brown; claspers with about distal two-thirds lightly sclerotised and semi-glossy, blackish with sclerotised part reddish brown to blackish; anal plate semi-glossy, pale reddish brown to blackish, edged with pale yellowish to reddish brown. Caudal horn of both green and dark forms semi-glossy; usually light reddish brown often with a suggestion of black apically or on some black larvae entirely blackish; 6.0–7.5 mm long; gently curved backwards; tapering evenly to a pointed apex; surface with scattered spine-like tubercles directed distally, blunt and very short, each with a simple seta, these tubercles similar in colour to adjacent part of horn. Length at maturity 85–100 mm. Width of head capsule 4.5–5.2 mm. Duration usually 6–9 days. Pupa (Pl. 8, figs i–k) Glossy; varying shades of brown, usually a little paler ventrally, without distinct markings, although often with some darkening primarily on head, metathorax and around anterior and posterior regions of abdomen. Proboscis with a trunk-like extension that has its distal third or so sharply recurved through 180º or more to adjoin or almost meet the ventral surface of thorax, the total length of the trunk-like extension, coloured usually dark brown to nearly black; proboscis along ventral midline brown, usually tipped black and reaching to apices of wings. Head, prothorax and mesothorax at x10 with surface finely rugose. Wings glossy, smooth, with veins barely raised; brown, sometimes edged black along distal margin. Antennae and legs brown often becoming blackish with maturity. Abdominal segments 1–8 with their anterior quarter finely pitted, otherwise almost smooth; abdominal segment 9 entirely rugose on dorsal and lateral surfaces, less so ventrally; spiracles distinct, black but lacking a surrounding black highlight; spiracular furrows with first ridge anterior of spiracle moderately developed, rarely with secondary ridges before cavity. Cremaster black, sometimes reddish brown basally; in dorsal view more or less an equilateral triangle, coarsely granulated; in lateral view tapering to a point; ventral surface similar in texture to dorsal surface, partially depressed; apex in dorsal view terminating in a pair of short, sharp spines in a V-shape. Length 42–48 mm. Width at widest point 11.3–12.5 mm. BIOLOGY Larval foodplants AMARANTHACEAE: Salsola sp. roly-poly, prickly saltwort. CONVOLVULACEAE: Convolvulus erubescens bindweed; 29/08/19 11:11:11.55 THE AUSTRALIAN FAUNA *Ipomoea batatas sweet potato, *I. indica morning glory, * Ipomoea sp. unidentified. Larvae from eggs laid by captured females fed on several species of Ipomoea, principally I. indica and I. batatas, the latter being preferred. It is often a very common species, sometimes occurring in huge numbers after heavy summer rains. There are records for all months except June and July, but adults probably occur throughout the year in northern Australia. Adults have been found feeding at dusk from lantana and Ixora blossom and the flowers of male pawpaws. There are no confirmed records of migration for A. godarti, but circumstantial evidence suggests migration occurs. Large aggregations of adults have been seen at Toowoomba on three separate occasions over some twenty years, all during February. On each occasion, the moths were present in numbers for only three or four nights suggesting they were migrating. Further, during continuous light trapping at Rockhampton in January 1983 and without having taken A. godarti in the preceding nights, Peter Mackey (pers. comm.) encountered five adults in a single night and 32 the next night but then none were seen for many weeks, again suggesting the moths were migrating. He monitored a light every night at Rockhampton for eight years with only one other similar incident; in February 1985, 17 A. godarti were attracted one night followed by five the following night, but no others were seen before or after for many weeks. A. wildei occurring also in New Guinea. Ambulyx dohertyi also extends its range into New Guinea (as A. dohertyi dohertyi) and eastward to New Britain, New Ireland and the Solomons as three additional subspecies (see species treatment). Adults are large with slender, pointed bodies and yellowish, brownish, or greyish wings, often highlighted with purple hues. The forewings are narrow with a costal margin that tends to curve distally to a falcate wing tip; the outer margin tends to be straight, mostly with a gently curved submarginal line or band, and most species have a large dark brown subbasal spot. The hindwings are often yellowish with suffused markings, the outer margin is usually crenulate and the anal angle is sharply defined. Male genitalia (Pl. 84, figs g, h) have an undivided uncus that tends to be apically bulbous and a short and often apically bifurcate gnathos; the harpe has either two well-developed spines or a dentate ridge often with a single spine; the saccus is distally slender; and the phallus has a long coecum and a dorsally protruding apex with two long, flat cornuti, sometimes with a third. Larvae are slender with numerous very small body tubercles and a long, slender, straight caudal horn, especially in early instars. In the 1st instar, the head is rounded but in the mid instars the vertex has bilobed protrusions developed to varying degrees, while in the last instar the vertex may be slightly conical or broadly rounded. Later instars are yellow to green, usually with a series of lateral oblique stripes of varying intensity. Although the immature stages of neither Australian species are known, it is anticipated that they would share similar characteristics with their known congeners. Foodplant records beyond Australia include members of the families Aceraceae, Anacardiaceae, Burseraceae, Dipterocarpaceae, Fagaceae and Juglandaceae (Robinson et al. 2001; Eitschberger and Ihle 2008, 2010). The families most commonly used by South-East Asian Ambulyx are Burseraceae and Anacardiaceae and as these two families are the only ones of the preceding list found in Australia, they may be among the foodplants for Australian Ambulyx species. Pupae are glossy brown and slender, the proboscis is confluent with the profile of the head and surrounding body and does not extend anterior of the head. The spiracular furrows are absent or ill-defined and the cremaster is triangular, terminating in two short spines. In the few examples in which the details of Ambulyx pupation have been reported, it occurs in a subterranean chamber. Kawahara et al. (2009) in their analysis of the higher classification of Sphingidae using five nuclear genes placed Ambulyx (represented by A. schauffelbergeri) sister to Amplypterus, which is widespread through the Indian subcontinent, eastward through China, South-East Asia to Timor with a single record from northern Australia. Genus Ambulyx Westwood, 1847 Ambulyx dohertyi queenslandi (Clark, 1928) Egg Laid singly on leaves or stems of the foodplant. The larvae, on emergence, usually eat the upper portion of the shell only. Larva Larvae are docile and are not overly responsive to disturbances. Larvae are usually found from November to May except in the tropical north where they tend to be associated with wet season rains. Early instar larvae rest on the undersides of juvenile leaves of the foodplant and feed from the leaf margin. Late instar larvae tend to rest on stems or under large leaves within lower sections of the foodplant and are remarkably well camouflaged. Pupation A mature larva placed in a container with 20 cm of soil burrowed to a depth of 17 cm, where it formed a cell with smoothed earthen walls. Parasitoids and predators TACHINIDAE: Blepharipa sp. (De Baar 2000). Adult Type species: Sphinx substrigilis Westwood, 1847. Designation by monotypy. Worldwide: 60 species, 20 subspecies (Kitching and Cadiou 2000; Brechlin 2005, 2006, 2009a, 2009b, 2014c; Eitschberger et al. 2006; Kobayashi et al. 2006; Brechlin and Kitching 2010a; Ivshin and Kitching 2014; Melichar and Řezáč [2014a]; Eitschberger 2015a; Melichar, Řezáč and Rindos 2015; Kitching et al. 2018b; Melichar et al. 2019). Australia: A. dohertyi queenslandi Clark, 1928; A. wildei Miskin, 1891. This large genus is widely spread across the Oriental region. The two Australian taxa are closely associated with the rainforests of the wet tropics of north-eastern Queensland with 190613 Hawkmoths of Australia 3pp.indd 65 65 Adults, Pl. 75; Pl. 84, fig. g. SYNONYMY Oxyambulyx dohertyi queenslandi Clark, 1928: 40 (type loc. Kuranda, Queensland). Ambulyx dohertyi dohertyi (Clark): Kitching and Cadiou, 2000: 79 (partim). Ambulyx dohertyi queenslandi (Clark): Brechlin and Kitching, 2010a: 21, 22, 23. Kitching and Cadiou (2000) synonymised the Australian subspecies, Ambulyx dohertyi queenslandi, with the nominotypical subspecies but retained subspecies solomonis (Rothschild and Jordan, 1903). Brechlin and Kitching (2010a) then reinstated subspecies queenslandi and described two additional subspecies, novobritannica from New 29/08/19 11:11:11.63 66 HAWKMOTHS OF AUSTRALIA Britain and novoirlandensis from New Ireland. More recently, Brechlin (2014c) added subspecies isabeliana from the Solomon Islands. DISTRIBUTION AND HABITAT Northern Queensland from Cape York to the Hervey Range near Townsville, at elevations from sea level to around 1100m (Mount Windsor Tableland). It is widespread through eastern Cape York Peninsula including from Lockerby Scrub, Jardine River headwaters, Moreton, Iron Range and the McIlwraith Range near Coen. Further south, there are a number of records from the Cooktown district both to the north and south. Most records are from the Wet Tropics, particularly the eastern escarpment of the Atherton Tablelands, where it can be found at localities such as Kuranda, Clohesy River and along the Palmerston Highway, and from the tropical coast between Daintree and Etty Bay to Tully, extending westwards along the Tully River valley. Records south of Tully are confined to the Paluma, Bluewater and Hervey Ranges (G. Cocks). Subspecies A. d. queenslandi is confined to Australia. Ambulyx dohertyi dohertyi is found in New Guinea. Other subspecies include A. d. novobritannica Brechlin and Kitching (2010a) from New Britain, A. d. novoirlandensis Brechlin and Kitching, 2010 from New Ireland, A. d. solomonis (Rothschild and Jordan, 1903) from Guadalcanal to Bougainville, Solomon Islands (Tennent 1999a), and A. d. isabeliana Brechlin, 2014 from Santa Isabel Island, Solomon Islands. ADULT DIAGNOSIS Male and female sexually dimorphic with differences in the colour and markings of the forewing above. Proboscis 21–26 mm long. Both sexes with a very dark green (tending black) longitudinal band between eyes continued along thorax passing by wing bases to distal margin, and abdomen with a very dark green lateral spot on segment 6, occasionally also on segment 7. Male forewings 41–47 mm long; above brown in varying shades and differing between individuals; a nearly circular, very dark green, subbasal spot rarely partly divided, a similar but much smaller spot on costal margin and another subtornal one; an inwardly curved dark brown line from apex fading before reaching tornus; discal spot absent or illdefined. Hindwings above mainly yellowish with three transverse scalloped bands, the inner two not always distinct; outer margin dark brown. Wings below shades of yellowish and reddish brown, the forewing with a distinct light grey outer margin. 190613 Hawkmoths of Australia 3pp.indd 66 Male genitalia (Pl. 84, fig. g) with uncus laterally flattened, broad and coming to a point at ventral apex; gnathos wide and deeply bifurcate with the two halves triangular, gradually tapering to a point; saccus triangular but apically narrowing; juxta semi-circular in outline with anterior margin straight; harpes with an inwardly projecting large, flat, triangular lobe at about midlength; phallus with a long, angled base and with a long, almost flat, apical projection dorsally about as long as phallus is wide, the vesica with three nearly flat bars, one long and blade-like with fine serrations on one side, the second of similar length but a little wider, less sclerotised and closely covered by short spines, the last one similar to the second but much smaller. Female forewings 46–53 mm long; above dark brown with a purplish sheen and markings mostly ill-defined, and the dark spots smaller than in the male and considerably variable in size between individuals; the pale discal spot usually distinct. Hindwings above, and fore- and hindwings below as in male. Ambulyx dohertyi queenslandi differs from A. wildei in having a dark lateral abdominal spot on segment 6 (sometimes also on 7), which is absent in A. wildei. Both A. d. queenslandi and A. wildei have a subbasal forewing spot, but A. d. queenslandi has an additional spot on the forewing costal margin. Further, the forewings of A. d. queenslandi females and some males have a purple sheen that is lacking in A. wildei, and the curved submarginal line is further from the outer margin in A. d. queenslandi than in A. wildei. DESCRIPTIONS OF IMMATURE STAGES Unknown. BIOLOGY Adults readily respond to light from late evening to the early morning with most being male, but females are not uncommon. To date, attempts to get a female to oviposit in captivity have been unsuccessful. Adults can be locally common and primarily fly during the wet season but can be found throughout the year at lower elevations. Ambulyx wildei Miskin, 1891 Adults, Pl. 75; Pl. 84, fig. h. SYNONYMY Ambulyx wildei Miskin, 1891: 20–21 (type loc. Cairns, Queensland). Oxyambulyx wildei (Miskin): Rothschild and Jordan, 1903: 204, pls 8, 30, 31. Oxyambulyx ceramensis Joicey and Talbot, 1921: 105 (type loc. Mount Manusela, Central Ceram, 6000 ft). Synonymised by Kitching et al. (2018b). Lectotype implicitly designated by D’Abrera [1987] as detailed by Kitching and Cadiou (2000). DISTRIBUTION AND HABITAT From northeastern Queensland south from Gap Creek near Mount Finlayson (south of Cooktown) to Tully and the Tully River valley. It is mainly a lowland species but is also recorded from the Windsor Tableland to 1000 m, and from Kuranda and the East Palmerston area up to 650 m. Adults can be locally common in the East Palmerston—Innisfail area. As the species also occurs in New Guinea, it is interesting that there are no Australian records from north of Cooktown despite extensive collections of hawkmoths from the rainforests of the McIlwraith Range, Iron Range and the far north of Cape York Peninsula. Its habitat is limited to coastal wet rainforest areas. 29/08/19 11:11:11.80 THE AUSTRALIAN FAUNA 67 Egg Lime green; glossy, smooth to the naked eye but under magnification (x50) with a finely rugose surface; ovoid, approximately 1.8 mm long x 1.7 mm wide x 1.7 mm high. BIOLOGY Adults fly primarily during the wet season, especially from February to March, but at lower elevations can be found throughout the year. They readily respond to light with females most often attracted in the late evening and males later, in the early morning hours. Unlike many sphingids in which the response to light is highly male biased, the ratio of females to males is approximately even. To date, several attempts to get females to oviposit in captivity have been unsuccessful. Genus Amplypterus Hübner, [1819] Beyond Australia, A. wildei is found in New Guinea. ADULT DIAGNOSIS Male and female sexually dimorphic with differences in the colour of the wings above. Proboscis 27–31 mm long. Both sexes with a very dark green (tending black) longitudinal band between eyes continuing along thorax passing over wing bases to distal margin. Male forewings 42–48 mm long; above greyish brown to brown in varying shades and differing between individuals; a nearly circular, very dark green, subbasal spot; an inwardly curved dark brown line from apex to tornus. Hindwings above mainly yellowish with three transverse dark bands, the inner one straight, the middle one scalloped, the outer one ill-defined; outer margin narrowly edged dark brown. Wings below shades of yellowish and reddish brown, the forewing with a distinct light grey outer margin. Male genitalia (Pl. 84, fig. h) with uncus laterally flattened, broad and coming to a point at ventral apex; gnathos short, broad and apically upturned and bifurcate; saccus subtriangular but apically narrowed; juxta triangular, laterally upturned and indented at midline on anterior margin; harpes with a large, flat, inwardly projecting, rounded-triangular lobe about midlength, bearing a robust inwardly curved spine on its margin; phallus with a very long apical projection ventrally that is nearly flat with a rounded apex, the vesica with two long, blade-like bars spined along one side, the most distal one slightly longer. Female forewings 48–54 mm long; above reddish brown, including the body, otherwise similar to male in maculation. Despite the large gap in distribution between A. wildei in Australia and in New Guinea there are no discernible differences in adult markings. Both A. wildei and A. dohertyi queenslandi have a subbasal forewing spot, but A. d. queenslandi has an additional spot on the costal margin. Ambulyx wildei also differs from A. d. queenslandi in having no abdominal spots, whereas A. d. queenslandi has a lateral spot on segment 6 and sometimes also on 7. Further, the forewings of A. wildei never have a purple sheen, which is present in females and some males of A. d. queenslandi, and the curved submarginal line is closer to the wing margin in A. wildei than in A. dohertyi queenslandi. DESCRIPTIONS OF IMMATURE STAGES Aside from eggs dissected from a fresh female, nothing is known of the immature stages. 190613 Hawkmoths of Australia 3pp.indd 67 Type species: Sphinx panopus Cramer, 1779. By subsequent designation by Kirby (1892) and discussed by Fletcher and Nye (1982). Worldwide: 2 species, 5 subspecies (Kitching and Cadiou, 2000; Eitschberger, 2006a; Melichar and Řezáč, [2014c]; Kitching et al. 2018b). Australia: A. panopus panopus (Rothschild and Jordan, 1903). Amplypterus species are distributed from India, Sri Lanka, the Andamans and China through South-East Asia to the Philippines, Indonesia, Timor and northern Australia. Adults are large, the forewings very long (more than twice the length of the hindwings), distinctly coloured light brown overlaid with varied darker brown shades, with an eyespot adjacent to the tornus (but not always distinct) and the outer margin weakly concave or straight just below the apex. The hindwings have the basal half diffused pink and the discal cell is small. The abdominal tergites are spined marginally, the tibiae are not spined and the eyes not lashed. The male genitalia have an undivided uncus that is slender and evenly curved, the gnathos is broad, short and apically bifurcate, the saccus is short, wide and broadly rounded; the juxta is broadly U-shaped and shorter than wide, the distally wide, broadly rounded valvae have a large dense patch of very short friction scales, the long slender harpes have an inwardly facing, flat, toothed projection at midlength, and the phallus is short with a large, downcurved coecum and a ventrally projecting apex bearing a small tooth ventrally. Larvae in the first instar have a rounded head and very long caudal horn. From the 2nd instar onward, the vertex of the head becomes conically elongated. Last instar larvae have oblique lateral stripes, a caudal horn that is exceedingly long and slightly curved forwards, the claspers are large and protrude distally beyond the anal plate, and the body carries numerous small tubercles. Larval foodplants include species of Anacardiaceae (including the commercial mango) and Clusiaceae. Pupae are semi-glossy, dark brown and thick-set, the proboscis is confluent with the profile of the head and surrounding body and does not extend anterior of the head. The spiracular furrows are well developed as four parallel ridges, and the cremaster is short, broad and triangular. Pupation occurs in a subterranean cell. In their analysis of the higher classification of Sphingidae using five nuclear genes Kawahara et al. (2009) found Amplypterus sister to Ambulyx. Amplypterus panopus panopus (Cramer, 1779) Common name: mango hawkmoth 29/08/19 11:11:11.96 68 HAWKMOTHS OF AUSTRALIA Adults, Pl. 75; Pl. 84, fig. i. SYNONYMY Sphinx panopus Cramer, 1779: 50, pl. 224, figs A, B (type loc. Jawa Tengah, Samarang [=Samarang, Java]). Calymnia panopus (Cramer): Walker, 1856: 124. Calymnia pavonica Moore, 1877: 596 (type loc. Port Blair, Andamanen). Amblypterus [sic] panopus (Cramer): Moore, 1882: 13. Amplypterus pavonicus (Moore): Kirby, 1892: 674. Compsogene panopus (Cramer): Rothschild and Jordan, 1903: 189. Compsogene (Calymnia) panopus (Cramer): Maxwell-Lefroy and Howlett, 1909: 467–468. Compsogene panopus panopus (Cramer): Rothschild and Jordan, 1907: 42. Amplypterus panopus panopus (Cramer): Bridges, 1993: VIII.3. Amplypterus panopus hainanensis Eitschberger, 2006a: 16-17, pl. 3, figs 5, 6 (type loc. Wuzhi Shan, 1500 m, Hainan, China). DISTRIBUTION AND HABITAT Only a single specimen is known from Australia, taken in Darwin, Northern Territory, in March 2009 by Paul Kay. The excellent condition of the specimen suggests it had recently emerged, but it is unclear whether it is a previously undetected resident, a recently established immigrant, or an occasional vagrant. Amplypterus panopus panopus is described as a woodland species in India (Bell and Scott 1937), and in Thailand it is found throughout the country up to 1700 m (Inoue, Kennett and Kitching 1997). Beyond Australia, A. panopus panopus occurs across the Indian subcontinent, eastward through China and SouthEast Asia to Alor Island in Indonesia (Eitschberger 2006a), and in Timor. There are four other subspecies: A. p. celebensis Rothschild and Jordan, 1903 from Sulawesi, A. p. karnatakaensis Melichar and Řezáč, [2014c] from southern India, A. p. mindanaoensis Inoue, 1996 from the Philippines, and A. p. sumbawanensis Eitschberger, 2006 from Flores and Sumba Islands, Indonesia (Kitching et al. 2018b). Two of these, A. p. celebensis and A. p. mindanaoensis, were given species status by Eitschberger (2006a) but returned as subspecies by Kitching et al. (2018b). Amplypterus panopus hainanensis Eitschberger, 2006 was synonymised with the nominotypical subspecies, and A. p. seramensis Inoue, 1999 from Seram was synonymised with subspecies A. p. celebensis by Kitching et al. (2018b). ADULT DIAGNOSIS Male and female similar but males are more darkly marked than females. Proboscis about 36 mm long. Forewings 55–80 mm long, with females usually larger than males; tending slender (about 2.5x longer than wide); above in many shades of brown with a straight blackish medial band paling on its distal margin and not quite parallel with outer margin; mainly whitish or very pale yellow distal of median band, becoming very pale pinkish brown towards apex, all irregularly spotted and a little variable between individuals; adjacent to tornus, a distinctive eyespot with a black and blurred grey centre finely encircled brown and with a fine semicircular edge of light blue. Hindwings above with basal half diffused pink and distal half in shades of brown, all overlaid with dark brown bands and lines of varying width, giving the species a distinctive appearance. Wings below mottled brown with forewing submarginal patch as above but more distinctly spotted. Male genitalia (Pl. 84, fig. i) with uncus undivided, evenly curved and gently tapering to a blunt point; gnathos short and broad, apically strongly upturned and deeply bifurcate; saccus short and wide, tapering to a broadly rounded apex; juxta broadly U-shaped, shorter than its diameter; valvae widest in distal half, broadly rounded with a large rounded dense patch of very short friction scales, sacculus slender, the harpe expanded into a flat, internally projecting triangular plate weakly toothed apically; phallus short, apex with a ventral projection with a small pimple ventrally, the coecum large, downcurved and about twothirds diameter of phallus; vesica tri-lobed. Amplypterus p. panopus differs from A. p. celebensis in lacking its yellowish submarginal forewing band that widens towards the apex and tornus. It differs from A. p. karnatakaensis in being a much paler moth, lacking the very black markings of A. p. karnatakaensis that dominate that subspecies. It differs from A. p. mindanaoensis in having less intense pink colouring on the hindwing. It differs from A. p. sumbawanensis in having the dark marking along the outer margin reduced to a very narrow linear edging before reaching the tornus, whereas in A. p. sumbawanensis the black marking is developed all the way to the tornus. DESCRIPTION OF IMMATURE STAGES Unknown from Australia. The life history has been documented by Mell (1922), Bell and Scott (1937), Dupont and Roepke (1941) and Eitschberger and Ihle (2008). The only images of the early instars are those of Eitschberger and Ihle (2008). Other images of late instar larvae and pupa are available on the internet. BIOLOGY Larval foodplants Beyond Australia, A. panopus panopus has been recorded on Dracontomelon dao, Mangifera caesia, M. indica mango, and Toxicodendron vernicifluum (all Anacardiaceae) and Calophyllum inophyllum and Garcinia oblongifolia (both Clusiaceae) (Robinson et al. 2001; Eitschberger and Ihle 2008). D’Abrera [1987] also lists Durio ‘durian’ (Malvaceae) but this requires confirmation. If A. panopus panopus is a resident in the Northern Territory, it may be using mango, the most common foodplant on the Indian subcontinent (Bell and Scott 1937). Among the other genera of recorded foodplants from South-East Asia, three genera, Toxicodendron, Calophyllum and Garcinia, also occur in the Northern Territory. Egg Eggs are laid singly on the underside of foodplant leaves. A captive female laid over 100 eggs on successive nights, the first of the eggs hatching after six days (Bell and Scott 1937). 190613 Hawkmoths of Australia 3pp.indd 68 29/08/19 11:11:12.14 282 HAWKMOTHS OF AUSTRALIA Plate 4. Acosmeryx anceus anceus: (a) egg; (b–i) larval instars as labelled [h, Eulophidae, Tetrastichinae, wasp larvae emerging from 3rd instar host; i, the same after parasitoid larvae had spun cocoons]; (j–l) pupae as labelled; (m) adult at rest. 190613 Hawkmoths of Australia 3pp.indd 282 29/08/19 11:11:37.98 PLATES 349 Plate 71. Theretra turneri: (a) egg; (b–h) larval instars as labelled; (i–k) pupae as labelled; (l) adult at rest. 190613 Hawkmoths of Australia 3pp.indd 349 29/08/19 11:12:07.95 PLATES 369 Plate 91. Male genitalia; spread with tegumen, uncus and gnathos turned to the right and phallus removed. (a) Tambo, Qld; (b) Grafton, NSW; (c) Edungalba, near Duaringa, Qld; (d) Forty Mile Scrub, N Qld; (e) Lizard Island, N Qld; (f) Tabubil, Papua New Guinea; (g) Forty Mile Scrub, N Qld; (h) Sumatra, Indonesia; (i) 80 km S of Larrimah, NT; (j) Dorrigo, NSW. 190613 Hawkmoths of Australia 3pp.indd 369 29/08/19 11:12:14.29 Appendix 1: Sphingidae–Parasitoid associations Sphingid host Parasitoid Acosmeryx anceus anceus Hymenoptera: Braconidae Cystomastax genus-group, probably a Macrostomion sp., det. DLJQ (ex host 5th instar larva, Clohesy River, near Kuranda, Qld) Hymenoptera: Eulophidae, subfamily Tetrastichinae, det. Stefan Schmidt) (ex host 3rd instar larva, Kuranda, Qld) Acosmeryx miskini Diptera: Tachinidae Blepharipa sp., det. BKC (ex host pupa, Brookfield, Qld) Agrius convolvuli Diptera: Tachinidae Blepharipa sp., det. BKC (Queensland Museum) Sturmia convergens (from Crosskey 1973) Agrius godarti Diptera: Tachinidae Blepharipa sp. (from De Baar 2000) Cephonodes janus Diptera: Tachinidae Blepharipa sp., det. BKC (ex host pupa, Rockhampton, Qld) Ceromya sp. 2, det. BKC (ex host larva, Rockhampton, Qld) Cephonodes kingii Diptera: Tachinidae Blepharipa sp., det. BKC (ex host pupa, Coen, Qld) Hymenoptera: Braconidae Microplitis basalis (from Austin & Dangerfield, 1993) Cizara ardeniae Diptera: Tachinidae Blepharipa sp., det. BKC (ex host pupa, Yungaburra, Qld), det. AR (ex host pupa, Avoca Beach, New South Wales) Winthemia sumatrana, det. BKC (ex host larva, Yungaburra, Qld) Coenotes eremophilae Diptera: Tachinidae Exorista psychidivora (from Cantrell, 1986) Palexorista sp. (from Cantrell 1986) Coequosa triangularis Hymenoptera: Eupelmidae Anastatus sp. 2, det. GAPG (ex host ovum, Wooyung NR, NSW) Hymenoptera: Ichneumonidae Lissopimpla excelsa det. GB (Dorrigo Plateau, NSW) Netelia sp., det. GB (Dorrigo Plateau, NSW) Daphnis moorei Hymenoptera: Scelionidae Telenomus sp. 1, det. LM (ex host ovum, Davies Creek, nr Kuranda, Qld) Hymenoptera: Trichogrammatidae Trichogramma sp., det. JMH (ex host ovum, Trinity Beach, Qld) Daphnis placida placida Diptera: Tachinidae Blepharipa sp., det. BKC (ex host pupa, Toowoomba, Qld) Ceromya sp., 2 det. BKC (ex host larva, Trinity Beach, Qld) Winthemia sumatrana det. BKC (ex host larva, Julatten, Qld) Hymenoptera: Scelionidae Telenomus sp. 1, det. LM (ex host ovum, Rockhampton, Qld) Hymenoptera: Eupelmidae Anastatus sp. 2, det. GAPG (ex host ovum, Rockhampton, Qld) Daphnis protrudens protrudens Hymenoptera: Scelionidae Telenomus sp. 1, det. LM (ex host ovum, Dinden State Forest, Qld) 190613 Hawkmoths of Australia 3pp.indd 372 29/08/19 11:12:14.60 Index Valid generic and species names for Australian hawkmoths, together with their primary page numbers, are in bold and generic names in capitals. abadonna, Sphinx 59 abbottii, Sphecodina 25 abdomen 6, 8, 10, 12, 14, 17 abelia 19, 62, 128 Abelia 93, 138, 148, 151, 179, 184, 185, 219 Abutilon oxycarpum 61 Acacia 272 farnesiana 98 Acanthaceae 94, 98, 376 Acari 29 accentifera, Nephele 205 accessory glands 12, 13 Aceraceae 65 acetosa, Ampelocissus 55, 260, 380 acetosa, Rumex 249 ACHERONTIA 6, 7, 10, 27, 45–47, 48, 58, 204 atropos 6, 23, 25, 27, 45, 46, 47, 48 atropos morta 47 australasiae 99 circe 47 lachesis 23, 27, 36, 39, 45, 46, 47–48 lachesis ab. radiata 47 lachesis diehli 47 lachesis f. fuscapex 47 lachesis f. pallida 47 lachesis f. submarginata 47 satanas 47 sojejimae 47 styx 27, 46, 47 Acherontiina 7, 38 Acherontiini 7, 27, 34, 47, 85, 151, 212 ACOSMERYX 11, 15, 48–49, 53 anceus 3, 48, 49, 52, 53, 56 anceus anceus 32, 35, 36, 40, 44, 48, 49–52, 53, 54, 55, 57, 372, 380 anceus alorana 49 anceus bismarckiana 49 anceus hainana 49 anceus halmaherana 49 anceus philippinensis 49 anceus subdentata 49 cinnamomea 3, 27, 35, 36, 40, 44, 45, 48, 49, 50, 51, 52–56, 57, 380, 381 daulis 49 miskini 27, 36, 40, 44, 48, 56–58, 372, 380, 381 mixtura 49 naga 49 acris, Cayratia 55, 128, 241, 266, 380 Actinidiaceae 122, 128, 251, 376 actinophylla, Alstonia 112, 115, 376 Acugutturidae 29 Acugutturus 29 aculeata, Pisonia 151, 378 acuminata, Vitex 223, 377 acuminatum, Santalum 98, 379 acutifolia, Plumeria 138, 376 Adansonia gibbosa 26 adfrontal suture 14 190613 Hawkmoths of Australia 3pp.indd 400 adnata, Cissus 55, 128, 249, 380 aedeagus 12 aeropyles 14, 15 aethiopica, Zantedeschia 136, 138, 260, 268, 376 African tulip tree 88, 228, 229, 232 africana africana, Kigelia 87, 376 africana, Olea 219, 378 Agaristinae 84 AGRIUS 7, 10, 15, 24, 26, 27, 58, 204 convolvuli 4, 6, 23, 24, 25, 26, 27, 30, 31, 36, 39, 41, 58–62, 63, 372, 377 convolvuli ab. fuscosignata 59 convolvuli ab. grisea 59 convolvuli ab. intermedia 59 convolvuli ab. major 59 convolvuli ab. minor 59 convolvuli ab. obscura 59 convolvuli ab. suﬀusa 59 convolvuli ab. unicolor 59 convolvuli ab. variegata 59 convolvuli ab. virgata 59 convolvuli var. ichangensis 59 convolvuli var. javanensis 59 convolvuli, var. tahitiensis 59 godarti 24, 26, 36, 39, 41, 58, 60 Agrotis infusa 29 Agryon 204, 374 Aidia racemosa 79, 81, 378 aksuensis, Herse convolvuli 59 Alangium villosum 115, 377 alata, Dillenia 128, 377 albata, Theretra nessus 254, 255 albibase, Macroglossum micacea 182, 183 albidus, Brachychiton 162, 380 Albizia basaltica 98, 377 albocitrinus, Coryceps 28 albolineata, Deilephila 135 alcedo, Macroglossa 167 alcedo, Macroglossum 11, 24, 36, 41, 43, 166, 167–169, 187, 188, 374, 379 alcicomis, Sarcorohdendorfia 128, 373 alecto, Theretra 23, 238 Aleuron biovatus 121 Alloxylon 84 Alocasia 118 brisbanensis 138, 260, 268, 376 cucullata 268, 376 macrorrhizos 138, 257, 260, 268, 376 alorana, Acosmeryx anceus 49 Alstonia 113, 275 actinophylla 112, 115, 376 constricta 112, 115, 376 muelleriana 112, 115, 376 scholaris 112, 115, 121, 376 althoferi, Prostanthera 98, 377 amara, Chaerocampa 247 amara, Theretra 247 Amaranthaceae 64, 94, 95, 98, 156, 159, 376 Amblypterus panopus 68 ambrymenis, Theretra insularis 246, 247 Ambulycini 10, 14, 34, 36, 108, 151 AMBULYX 10, 65, 67 dohertyi 65 dohertyi dohertyi 65, 66 dohertyi isabeliana 66 dohertyi novobritannia 65 dohertyi novoirlandensis 66 dohertyi queenslandi 36, 65–66, 67 dohertyi solomonis 65, 66 schauﬀelbergeri 65 wildei 36, 65, 66–67 americanus, Gyrocarpus 96, 98, 213, 216 Amorphophallus bulbifer 249, 376 Amorphophallus paeoniifolius 260, 376 Ampelocissus acetosa 55, 260, 380 frutescens 128, 138, 246, 260, 380 gardineri 138, 260, 380 Amphimermis bogongae 29 Amphimoea walkeri 6 Amphion, brennus 132 Amphion, floridensis 25 AMPLYPTERUS 10, 65, 67 panopus 68 panopus celebensis 68 panopus hainanensis 68 panopus karnatakaensis 68 panopus mindanaoensis 68 panopus panopus 35, 36, 67–69 panopus seramensis 68 panopus sumbawanensis 68 pavonicus 68 ampulla 10 Anacardiaceae 65, 67, 68 anal angle 7 anal lobe 7 anal plate 15, 18 anal prolegs 15 anal tube 10 anal vein 7, 9 Anastatus 31, 107, 115, 125, 372, 373, 374 bifasciatus 31 biproruli 31 pearsalli 31 anceus, Acosmeryx 3, 48, 49, 52, 53, 56 anceus, Enyo 49 anceus, Sphinx 49 anceus alorana, Acosmeryx 49 anceus anceus, Acosmeryx 32, 35, 36, 40, 44, 48, 49–52, 53, 54, 55, 57, 372, 380 anceus bismarckiana, Acosmeryx 49 anceus hainana, Acosmeryx 49 anceus halmaherana, Acosmeryx 49 anceus philippinensis, Acosmeryx 49 anceus subdentata, Acosmeryx 49 anchemolus, Eumorpha 122 andamana, Daphnis 112 andreana, Saurauia 128, 376 andronical tufts 10, 122 anellus 11, 12 ANGONYX 22, 34, 69, 210, 211 emilia 69 excellens 211 papuana 69, 377 papuana bismarcki 69 papuana papuana 36, 40, 45, 69–71, 211 papuana papuana f. serrata 69 serrata 69 testacea 69 29/08/19 11:12:18.97 INDEX testacea papuana 69 Angophora 102 costata 102, 377 Angraecum sesquipedale 26 angustans, Choerocampa 112 angustans, Daphnis 112 angustifolia, Fraxinus 219, 378 angustilobum, Typhonium 260, 376 angustisepala, Ervatamia 115 angustissima, Clematicissus 138, 380 anne, Psilogramma 226 Annona muricata 87 ant plant 25, 93, 175 antarctica, Cissus 52, 55, 58, 128, 241, 243, 244, 249, 380 anteclypeus 14 antennae 6, 7, 14, 16, 26, 17 anterior apophysis 13, 14 Anthocephalus chinensis 112 Anthurium plowmanii 120, 376 Anthurium schlechtendalii 120, 376 antipoda, Zonilia 208 Antirrhinum majus 219, 378 antrum 13 ants 25, 29, 175 bull ants (Myrmecia) 29 green tree ants (Oecophylla) 29 myrmecodia ants (Philidris) 175 Anumara 27 Aphelinidae 31 Apis 6, 27, 46, 47 cerana 46, 47 dorsata 27, 46, 47 koschevnikovi 46 mellifera 27, 46, 47 Apocynaceae 75, 94, 98, 109, 112, 115, 121, 122, 129, 138, 205, 208, 210, 235, 275, 237, 376 apollo, Hypochrysops 175 apophysis 13, 14 approximans, Macroglossa 169 approximans, Macroglossum corythus 3, 25, 35, 36, 40, 43, 166, 169–172, 173, 179, 183, 374, 378, 379 approximata, Macroglossa 201 aquatica, Ipomoea 61, 377 aquila, Theretra 263 arabica, Coﬀea 79 Araceae 58, 118, 120, 129, 136, 138, 238, 249, 251, 257, 260, 266, 267, 268, 376 Araliaceae 96, 98, 376 arcuatum, Macroglossum 172 ardenia, Deilephila 91 ardenia, Sphinx 91 ardenia, Zonilia 91 ardeniae, Cizara 4, 8, 27, 36, 39, 45, 91–93, 201, 372, 378, 379 ardeniae, Sphinx 91 argentata, Chaerocampa 257 argentata, Deilephila 257 argentata, Sphinx 257 argenteus, Pipturus 240, 241, 263, 380 argos, Psilogramma 11, 32, 35, 36, 39, 41, 211, 212, 213–216, 375, 377 arida, Coenotes 5, 35, 36, 40, 42, 94–95, 96, 376 arolium 8, 10 arrhenotokous parthenogenesis 30 artificial diets 23 aruensis, Psilogramma mastrigti 229 ash claret 219 golden 219 Himalayan 219 velvet 219 Asian bell tree 219 190613 Hawkmoths of Australia 3pp.indd 401 Asperula 148 conferta 147, 378 Asteraceae 58, 147, 148 Asystasia gangetica 189 Atractocarpus fitzalanii 79, 81, 378 Atractocarpus sessilis 81, 378 atropivora, Zygobothria 219, 375 Atropos 45 atropos morta, Acherontia 47 atropos, Acherontia 6, 23, 25, 27, 45, 46, 47, 48 atropos, Sphinx 45 attenuata, Canthium 84, 273 attenuata, Psydrax 84, 273, 379 augusta, Gardenia 75, 79, 84 aureum 120 auricularia, Exallage 132 auricularia, Oldenlandia 132 australasiae, Acherontia 99 australasiae, Brachyglossa 99 australasiae, Coequosa 5, 15, 23, 24, 32, 36, 37, 43, 99–103, 377 australasiae, Metamimas 98, 99 australasiae, Phryxus livornica, var. 157 australasiae, Sphinx 98, 99 australasica, Raphidophora 120, 376 australiensis, Gnathothlibus 5, 35, 36, 37, 44, 45, 122–125, 126, 127, 128, 373, 377 australiensis, Pavetta 75, 79, 178, 379 australis, Cephonodes 3, 11, 27, 31, 35, 36, 40, 71, 72–76, 77, 78, 80, 81, 83, 84, 379 australis, Cephonodes hylas 72 australis, Emex 138, 378 austrosundanus, Cephonodes janus, 79 babarensis, Theretra celata 239 Bacillus thuringiensis (BT) 28 backi, Macroglossum 201 backi, Macroglossum vacillans 201 bacteria 28 balsam 27, 138, 147, 249, 260 balsamina, Impatiens 138, 147, 249, 260 Balsaminaceae 129, 138, 147, 238, 249, 251, 260, 376 bandicoot berry 128, 243, 249 Banksia ericifolia 106, 378 integrifolia 106, 378 marginata 106, 378 serrata 106, 378 spinulosa 106, 378 banksiae, Brachyglossa 99 banksiae, Metamimas 99 Barleria cristata 98, 376 basalis, Microplitis 84, 260, 372, 375 basaltica, Albizia 98, 377 basitarsus 8, 10 batatas, Ipomoea 61, 65, 128, 138, 148, 260, 377 beach gardenia 172, 173 Beauveria 28, 138 beccarii, Myrmecodia 93, 175, 379 beddoesii, Hippotion 148 bedstraw 147 bee hawkmoths 71 Begonia (begonia) 138, 251, 376 Begoniaceae 138, 251, 376 belinda, Macroglossa 176 bernardus, Chaerocampa 145 berteroana, Fagraea 232, 377 bethia, Diludia 163 bethia, Leucomonia 5, 36, 39, 41, 163–166, 374, 377 bethia, Macrosila 163 401 bethia, Meganoton 163 Betulaceae 108, 277 bhaga, Eurypteryx 121 bicolor, Euplectrus 31 bifasciatus, Anastatus 31 bignonia 219 Bignoniaceae 85, 87, 129, 138, 205, 212, 219, 228, 232, 238, 260, 376–377 biguttata, Hyles 157 bilineata, Clanis 27 billardierianum, Epilobium 147, 378 bindweed 64 biovatus, Aleuron 121 biproruli, Anastatus 31 bird-catcher tree 151 bird’s nest anthurium 120 bismarcki, Angonyx 69 bismarcki, Angonyx papuana 69 bismarcki, Theretra indistincta 242 bismarckiana, Acosmeryx anceus 49 Blepharipa 33, 58, 62, 65, 81, 84, 93, 115, 120, 128, 138, 141, 163, 189, 219, 250, 257, 260, 266, 269, 372, 373, 374, 375 fulviventris 138, 257, 373, 375 Blondeliini 33 Boerhavia aculeata 151, 378 chinensis 138, 159, 377 diﬀusa 138, 159, 377 dominii 138, 254, 378 pubescens 159, 254, 378 boerhaviae, Chaerocampa 129 boerhaviae, Hippotion 37, 38, 44, 128, 129–132, 142, 143, 373, 377 boerhaviae, Sphinx 129 bogongae, Amphimermis 29 Bombylia 166 Bombyliinae 34 Borreria exserta 145, 378 Bougainvillea spectabilis 151, 378 bowmanii, Eremophila 98, 379 brachycera, Cosmotriche 151, 152 brachycera, Hopliocnema 15, 36, 37, 43, 89, 90, 91, 151, 152–154, 155, 156, 373, 379 Brachychiton 160, 162 albidus 162, 380 chillagoensis 162, 380 paradoxus 162, 380 Brachyglossa, australasiae 99 Brachyglossa, banksiae 99 Brachymeria 31, 128, 204, 373, 374 Braconidae 31, 32, 52, 84, 181, 203, 216, 260, 372, 375 brasiliensis, Richardia 145, 260, 379 brennus, Amphion 132 brennus, Chaerocampa 132 brennus, Hippotion 3, 35, 37, 38, 44, 128, 129, 132–135, 138, 139, 189, 373, 377, 378, 379 brennus, Sphinx 132 brennus, Theretra 132 brennus f. brennus, Hippotion 132 brennus f. funebris, Hippotion 132 brennus f. johanna, Hippotion 138 brennus f. rubribrenna, Hippotion 132, 133 brennus f. viettei, Hippotion 133 brennus funebris, Hippotion 132, 133 brennus johanna, Hippotion 138 brennus viettei, Hippotion 132 Breonia 275 Breonia chinensis 112, 378 brisbanensis, Alocasia 138, 260, 268, 376 brownii, Pavetta 75, 379 brownii, Typhonium 138, 376 29/08/19 11:12:19.09 402 HAWKMOTHS OF AUSTRALIA brycei, Podranea 138, 376 bucklandii, Cephonodes 82 bucklandii, Hemaris 82 Buddleia 93, 184 bulbifer, Amorphophallus 249, 376 bulbifera, Dioscorea 257, 271, 377 burica, Eupanacra splendens 119 bursa copulatrix 13 Burseraceae 65 Buxaceae 277 Caesalpiniaceae 84 caesia, Mangifera 68 cairica, Ipomoea 61, 377 Caladium 118 caleyi, Grevillea 106, 378 Calliandra riparia 189 callistegioides, Clytostoma 219, 260, 376 Callosphingia 58, 204 callusia, Daphnis dohertyi 109, 110 Calophyllum 68 inophyllum 68 Calosoma schayeri 95, 159, 160 Calosotinae 31 Calymnia panopus 68 Calymnia pavonica 68 Calystegia sepium 61 calyx 13 camara, Lantana 75, 79, 115, 179, 189 cambagei, Fagraea 232, 377 camouflage 25, 27 campanulata, Spathodea 88, 205, 219, 228, 229, 232, 377 Campsis grandiflora 219, 376 Campsis radicans 219, 376 Cananga odorata 87, 376 Candollea serrulata 138 Canthium 184 attenuata 84, 273 coprosmoides 84 odorata 75, 79, 81, 84, 178 oleifolia 84 ridigula 273 cape honeysuckle 219 capensis, Tecomaria 219, 377 capensis, Theretra 238 Caprifoliaceae 212, 219, 277, 377 Caquosa 98 carandas, Carissa 206, 208 Carcelia 33, 120, 269, 373, 374, 375 hackeri 374 kockiana 193 prominens 159, 373 Carceliini 33 cardiophylla, Cayratia 243, 244, 380 careya, Planchonia 260, 377 Carica papaya 26, 79, 138, 169, 179, 184, 189, 260, 266 Carissa 205, 206 carandas 206, 208 edulis 208 lanceolata 98, 210, 376 laxiflora 208, 210, 376 ovata 210, 376 spinarum 208 Castanopis 109 Casuarina 219 Casuarinaceae 219 casuarinae, Diludia 216 casuarinae, Macrosila 216 casuarinae, Meganoton 216 casuarinae, Psilogramma 17, 25, 27, 35, 36, 39, 41, 166, 211, 212, 213, 216–219, 221, 226, 229, 375, 376, 377, 378, 380 casuarinae, Sphinx 216 190613 Hawkmoths of Australia 3pp.indd 402 caudal horn 15, 16, 18 cavicola, Hexamermis 29 cayennensis, Stachytarpheta 26, 98, 151, 179, 184, 189, 193, 380 Cayratia 49, 251, 258 acris 55, 128, 241, 266, 380 cardiophylla 243, 244, 380 clematidea 25, 52, 55, 58, 128, 138, 145, 147, 241, 243, 244, 249, 260, 266, 380 maritima 249, 380 trifolia 128, 260, 271, 380 Cechenena 12, 238 Cechetra 238, 246 Celastraceae 277 celata, Chaerocampa 239 celata, Theretra 35, 239 celata, Theretra clotho 239 celata babarensis, Theretra 239 celata celata, Theretra 37, 38, 44, 238, 239–241, 242, 380 celebensis, Amplypterus panopus 68 celerio, Choerocampa 135 celerio, Chaerocampa (Theretra) 135 celerio, Deilephila 135 celerio, Hippotion 4, 23, 27, 28, 33, 37, 44, 128, 129, 135–138, 159, 160, 245, 373, 376, 377, 378, 379, 380, 381 celerio, Hippotion (Chaerocampa) 135 celerio, Sphinx 128, 135 celerio, Theretra 135 celerio ab. brunnea, Hippotion 135 celerio ab. pallida, Hippotion 135 celerio ab. unicolor, Hippotion 135 celerio f. luecki, Hippotion 136 celerio f. rosea, Hippotion 136 celerio f. sieberti, Hippotion 135 Celerio lineata livornicoides 157 celerio var. augustii, Deilephila 135 Centrodora darwini 31 CEPHONODES 6, 7, 10, 11, 15, 19, 22, 25, 26, 37, 45, 71–72, 76, 80, 81, 82, 83 australis 3, 11, 27, 31, 35, 36, 40, 71, 72–76, 77, 78, 80, 81, 83, 84, 379 bucklandii 82 cunninghami 3, 7, 27, 35, 36, 40, 71, 72, 73, 74, 75, 76–79, 80, 81, 83, 84, 378, 379 hylas 7, 23, 24, 25, 27, 31, 35, 71, 72, 73, 76, 77 hylas australis 3, 72 hylas cunninghami 72, 76 hylas melanogaster 72 hylas virescens 72 janus 22, 36, 40, 71, 73, 75, 77, 78, 79–82, 83, 84, 372, 378, 379 janus austrosundanus 79 janus simplex 79 kingi 82 kingii 16, 24, 27, 32, 36, 40, 62, 71, 73, 75, 77, 78, 80, 81, 82–84, 201, 274, 372, 378, 379 picus 3, 35, 71, 72, 73, 76, 77 unicolor 79 xanthus 72 cerana, Apis 46, 47 Ceratopogonidae 30 CERBERONOTON 5, 10, 11, 26, 34, 84–85 loeﬄeri 85 rubescens 3, 35, 85, 86 rubescens philippinensis 86 rubescens rubescens 85, 86 rubescens severina 34, 85 rubescens thielei 85, 86 rubescens titan 86 severina 3, 24, 34, 35, 36, 39, 41, 58, 84, 85–88, 104, 376, 377 Ceromya 33, 81, 115, 125, 250, 372, 373, 375 Chaerocampa, see also Choerocampa amara 247 argentata 257 bernardus 145 boerhaviae 129 brennus 132 celata 239 celerio 135 cleopatra 241 cloacina 239 comminuens 247 curvilinea 241 deserta 247 drancus 257 eras 125 erotus var. eras 125 firmata 257 ignea 145 inornata 244 insularis 246 intersecta 263 johanna 138 latreillii 247 lucasii 250 luteotincta 239 margarita 251 marginata 251 nessus 254 nessus var. rubicundus 254 oldenlandiae 257 pallicosta 4 pallida 244 phoenix 251 potentia 260 procne 250 puellaris 257 queenslandi 260 rosetta 141 sapor 125 scrofa 145 sobria 257 swinhoei 148 tenebrosa 250 tryoni 266 velox 148 walduckii 247 Chalcididae 31 Chalcidoidea 31–32 Charletonia 29 charon, Spectrum 47 CHELACNEMA 3, 5, 7, 14, 22, 24, 26, 34, 88, 151, 152, 233 ochra 3, 5, 8, 24, 25, 34, 36, 40, 43, 88–91, 151, 152, 154, 155, 379, 380 cheni, Macroglossum ungues 276 chillagoensis, Brachychiton 162, 380 Chinchona 275 chinensis, Anthocephalus 112 chinensis, Boerhavia 138, 159, 377 chinensis, Breonia 112, 378 Chionanthus ramiflorus 228, 378 chiron, Nephele 205 chiron, Sphinx 205 chiron, Xylophanes 205 chiron, Zonilia 205 chlorostachys, Erythrophleum 84 Choerocampa, see also Chaerocampa angustans 112 celerio 135 equestris 254 hesperus 112 indistincta 241 29/08/19 11:12:19.23 INDEX neriastri 115 pallicosta 4 procne 250 protrudens 115 yorkii 148 Choerocampina 7, 11, 34, 37, 118, 122 chorion 14, 15 choui, Psilogramma 220 Chromis erotus cramptoni 125 Chromis erotus eras 125 cingulata, Agrius 58 cingulata, Sphinx 58 cinnamomea, Acosmeryx 3, 27, 35, 36, 40, 44, 45, 48, 49, 50, 51, 52–56, 57, 380, 381 cinnamomea, Enyo 52, 53 circe, Acherontia 47 Cissus 49, 84, 93, 251 adnata 55, 128, 249, 380 antarctica 52, 55, 58, 128, 241, 243, 244, 249, 380 hypoglauca 128, 380 oblonga 55, 58, 128, 147, 241, 246, 249, 260, 380 penninervis 128, 380 reniformis 243, 246, 380 repens 128, 241, 249, 380 rhombifolia 128, 249, 380 Citharexylum hidalguense 219, 380 citrifolia, Morinda 22, 128, 167, 172, 173, 178, 186, 192, 193, 201, 379 citriodora, Corymbia 102, 377 Citrus 251 limon 84 CIZARA 7, 22, 91 ardeniae 4, 8, 27, 36, 39, 45, 91–93, 201, 372, 378, 379 sculpta 91 Clanis bilineata 27 Clarkia concinna 260, 378 unguiculata 260, 378 claspers 15, 18 Clematicissus angustissima 138, 380 opaca 128, 138, 246, 249, 260, 380 clematidea, Cayratia 25, 52, 55, 58, 128, 138, 145, 147, 241, 243, 244, 249, 260, 266, 380 cleopatra, Chaerocampa 241 cleopatra, Theretra 241 Clerodendrum 48, 61, 166 floribundum 61, 98, 163, 165, 219, 223, 225, 232, 377 paniculatum 232, 377 tomentosum 26, 219, 377 tracyanum 232, 377 cloacina, Chaerocampa 239 cloacina, Theretra 239 clotho, Theretra 239, 240, 241 clotho celata, Theretra 239 clotho manuselensis, Theretra 3, 241 clotho papuensis, Theretra 3, 241 Clusiaceae 67, 68 clypeus (of larva) 14, 16 Clytostoma callistegioides 219, 260, 376 coccinea, Ixora 115 cocky apple 260 cocytoides, Meganoton 85 coecum 11, 12 Coelonia 7, 47, 58, 204 Coelospermum 93 paniculatum var. syncarpum 93, 186, 201, 378 reticulatum 135, 172, 178, 179, 260, 378 190613 Hawkmoths of Australia 3pp.indd 403 COENOTES 24, 34, 93–94, 160 arida 5, 35, 36, 40, 42, 94–95, 96, 376 eremophilae 25, 27, 36, 40, 42, 94, 95–98, 159, 372, 376, 377, 378, 379, 380 COEQUOSA 6, 14, 15, 22, 24, 25, 98–99, 102 australasiae 5, 15, 23, 24, 32, 36, 37, 43, 99–103, 104, 377 triangularis 7, 15, 23, 24, 27, 32, 36, 37, 43, 98, 99, 102, 103–108, 372, 378 Coﬀea arabica 79 coffee 79 coffin flies (Phoridae) 33 colliculum 13 Colocasia 251 esculenta 138, 260, 268, 376 colour morphs (larva) 25 comminuens, Chaerocampa 247 common oviduct 13 compound eyes 6, 8 Compsilura concinnata 33, 138, 373 Compsogene (Calymnia) panopus 68 Compsogene panopus 68 concinna, Clarkia 260, 378 concinnata, Compsilura 33, 138, 373 conferta, Asperula 147, 378 constricta, Alstonia 112, 115, 376 convergens, Sturmia 62, 372 Convolvulaceae 58, 59, 61, 64, 128, 129, 138, 148, 260, 377 convolvuli, Agrius 4, 6, 23, 24, 25, 26, 27, 30, 31, 36, 39, 41, 58–62, 63, 372, 377 convolvuli, Macrosila 59 convolvuli, Phlegethontius 59 convolvuli, Protoparce 59 convolvuli, Sphinx 58, 59 convolvuli ab. alicea, Sphinx 59 convolvuli ab. extincta, Herse 59 convolvuli ab. fasciata, Protoparce 59 convolvuli ab. fuscosignata, Agrius 59 convolvuli ab. grisea, Agrius 59 convolvuli ab. intermedia, Agrius 59 convolvuli ab. major, Agrius 59 convolvuli ab. minor, Agrius 59 convolvuli ab. obscura, Agrius 59 convolvuli ab. suﬀusa, Agrius 59 convolvuli ab. unicolor, Agrius 59 convolvuli ab. variegata, Agrius 59 convolvuli ab. virgata, Agrius 59 convolvuli aksuensis, Herse 59 convolvuli f. posticoconflua, Herse 59 convolvuli indica, Protoparce 59 convolvuli marshallensis, Herse 59 convolvuli peitaihoensis, Herse 59 convolvuli roseafasciata, Sphinx 59 convolvuli var. batatae, Sphinx 59 convolvuli var. distans, Sphinx 59 convolvuli var. ichangensis, Agrius 59 convolvuli var. javanensis, Agrius 59 convolvuli var. nigricans, Sphinx 59 convolvuli var. tahitiensis, Agrius 59 Convolvulus 61, 260 erubescens 64, 377 convolvulus hawkmoth 58 Coprosma 91 lucida 93, 378 quadrifida 93, 378 repens 93, 147, 178, 378 coprosmoides, Canthium 84 coprosmoides, Cyclophyllum 84, 378 cordata, Philidris 175 Cordyceps 28 403 Cordycipitaceae 28 corn plant 120, 184, 204 Cornaceae 109, 115, 377 cornuti (cornutus) 12 coronal suture 14, 16 corpus bursae 13, 14, 21 Corymbia citriodora 102, 377 corythus, Macroglossa 172 corythus, Macroglossum 169, 170, 171 corythus approximans, Macroglossum 3, 25, 35, 36, 40, 43, 166, 169–172, 173, 179, 183, 374, 378, 379 corythus corythus, Macroglossum 36, 166, 170, 172–173, 379 corythus fulvicaudata, Macroglossum 170 corythus pylene, Macroglossum 3, 169, 170 Cosmotriche brachycera 151, 152 costata, Angophora 102, 377 Cotoneaster 219 countershading 29 coxa 7, 8, 16 crameri, Daphnis hypothous 35, 36, 109, 275 cramptoni, Chromis erotus 125 cremaster 17 crepe myrtle 249 Crinum pedunculatum 26 cristata, Barleria 98, 376 crochets 15, 16 cucullata, Alocasia 268, 376 cunninghami, Cephonodes 3, 7, 27, 35, 36, 40, 71, 72, 73, 74, 75, 76–79, 80, 81, 83, 84, 378, 379 cunninghami, Cephonodes hylas 72, 76 cunninghami, Hemaris 76 cunninghami, Macroglossa 76, 79 cunninghami, Sesia 76 Curculigo ensifolia 132 377 currant bush 210 curvilinea, Chaerocampa 241 curvilinea, Theretra 241 cyanoides, Melastoma 125, 377 Cyclophyllum coprosmoides 84, 378 CYPA 6, 8, 10, 108 decolor 108, 109 decolor decolor 108, 109 decolor euroa 35, 36, 108–109 ferruginea 108 uniformis 109 Cyrtosperma johnstonii 268, 376 Cystomastax 52, 372 Dahlia 147 dalii, Deilephila 208 dallachiana, Tarenna 79, 379 dalrympleana, Gmelina 232, 377 danneri, Psilogramma 220 dao, Dracontomelon 68 DAPHNIS 11, 14, 109, 110, 111, 116, 167, 205 andamana 112 angustans 112 dohertyi 15, 109 dohertyi dohertyi 36, 109–110 dohertyi callusia 109, 110 gigantea 110 gloriosa 110 hesperus 112 horsfieldii 112 hypothous crameri 35, 36, 109, 275 hypothous moorei 110 hypothous pallescens 110 jamdenae 113 magnifica 110 29/08/19 11:12:19.37 404 HAWKMOTHS OF AUSTRALIA moorei 36, 37, 43, 45, 109, 110–112, 113, 116, 275, 372, 378, 379 nerii 23, 30, 109 pallescens 110 placida 112 placida placida 36, 40, 43, 45, 109, 111, 112–115, 116, 372, 376, 377 placida salomonis 113 protrudens 111, 113, 115, 116, 379 protrudens lecourti 116 protrudens protrudens 36, 37, 40, 43, 45, 109, 115–118, 372 torenia rosacea 112 daphnoides, Psychotria 178, 379 Daphnusa 277 miskini 56 Darapsa, eras 125 Darapsa, moorei 4, 110 Darapsa, placida 112 darius, Macrosila 219 darlingtoni, Panacra excellens 211 darwini, Centrodora 31 daulis, Acosmeryx 49 death’s head hawkmoths 27, 45, 46 decipiens, Persicaria 26, 141, 378 decolor, Cypa 108, 109 decolor decolor, Cypa 108, 109 decolor euroa, Cypa 35, 36, 108–109 Deidamia 49 Deilephila 238 albolineata 135 ardenia 91 argentata 257 celerio 135 celerio var. augustii 135 dalii 208 dohertyi 109 elpenor 25, 26, 27 eras 125 gigantea 110 jamdenae 113 livornica 157 livornicoides 157 oldenlandiae 257 pallescens 110 placida placida 113 porcellus 4 porcia 145 proxima 257 protrudens 115 scrofa 145 spilota 250 deliciosa, Monstera 120, 376 Dendrocnide 25, 240 excelsa 241, 263, 380 moroides 263, 380 photinophylla 241, 263, 380 depictum, Hippotion 129, 141, 142 Deplanchea tetraphylla 232, 376 deserta, Chaerocampa 247 deserta, Theretra 247 deserti, Eremophila 98, 379 devil’s ivy 120 diaphragm 11, 12 didyma, Nephele 205 didyma, Sphinx 205 didyma ab. hespera, Nephele 205 didyma f. didyma, Nephele 205 didyma f. hespera, Nephele 205 didymum, Jasminum 223, 378 Dieﬀenbachia 118 diehli, Acherontia lachesis 47 diﬀusa, Boerhavia 138, 159, 377 Dillenia alata 128, 377 Dilleniaceae 69, 122, 128, 129, 135, 138, 238, 260, 377 190613 Hawkmoths of Australia 3pp.indd 404 Dilophonotini 15, 34, 72 Diludia bethia 163 casuarinae 216 godarti 62 latreillii 247 macromera 220 melanomera 220 nebulosa 226 obliqua 204 rubescens 84 Dioscorea bulbifera 257, 271, 377 discolor 257 dodecaneura 257 transversa 271, 377 Dioscoreaceae 238, 257, 271, 377 Dipterocarpaceae 65, 108, 109 Dipterocarpus lanceolata 109 tuberculatus 109 discal cell 7, 9 discistriga, Macrosila 219, 220 discistriga discistriga, Psilogramma 35, 36, 211, 212, 219–220, 227 discistriga hayati, Psilogramma 220 discocellular crossvein 7, 9 discolor, Dioscorea 257 dissecta, Merremia 61, 377 dissecting genitalia 20–21 distans, Protoparce 59 distans var., Sphinx convolvuli 59 distincta, Phlegethontius 62 distincta, Sphinx 62 distincta, Theretra latreilleii 250 distincta f., Theretra latreillei lucasi 250 distinctum, Meganoton 163 divaricata, Tabernaemontana 115, 376 divergens, Macroglossum 3, 193 divergens queenslandi, Macroglossum 193 divergens, Macroglossum heliophila 193 doddi, Macroglossum 173 doddi, Macroglossum dohertyi 25, 26, 36, 39, 43, 44, 92, 166, 169, 173–176, 374, 379 dodecaneura, Dioscorea 257 dohertyi, Ambulyx 65 dohertyi, Daphnis 15, 109 dohertyi, Deilephila 109 dohertyi, Macroglossum 167, 175 dohertyi, Panacra 118 dohertyi callusia, Daphnis 109, 110 dohertyi doddi, Macroglossum 25, 26, 36, 39, 43, 44, 92, 166, 169, 173–176, 374, 379 dohertyi dohertyi, Ambulyx 65 dohertyi dohertyi, Daphnis 36, 109–110 dohertyi dohertyi, Macroglossum 173, 174 dohertyi isabeliana, Ambulyx 66 dohertyi melanura, Macroglossum 173 dohertyi novobritannia, Ambulyx 65, 66 dohertyi novoirlandensis, Ambulyx 66 dohertyi queenslandi, Ambulyx 36, 65–66, 67 dohertyi queenslandi, Oxyambulyx 65 dohertyi solomonis, Ambulyx 65, 66 dominii, Boerhavia 138, 254, 378 dorsata, Apis, 27, 46, 47 double-headed hawkmoth 103 Dracaena fragans 120, 148, 169, 184, 204 Dracontomelon dao 68 drancus, Chaerocampa 257 drancus, Sphinx 257 drancus, Xylophanes 257 Drino 33, 263, 375 Duboisia 98 leichhardtii 98, 380 myoporoides 98, 380 ductus bursae 13, 14 ductus ejaculatorius 12 ductus seminalis 13 dumolinii, Lophostethus 27 Duranta (duranta) 19, 75, 79, 151, 184 repens 82, 84, 219, 380 dyars, Smerinthus 276 dyras tenimberi, Marumba 277 ectoparasitoids 30, 31, 32 edulis, Carissa 208 edwardsi, Macrosila 235 edwardsi, Meganoton 235 edwardsi, Psilogramma 235 edwardsi, Tetrachroa 11, 36, 39, 45, 235–238, 376 Ehretiaceae 58 Elaeagnaceae 156 elata, Eucalyptus 102, 377 elliptica, Ochrosia 115, 376 Elpenor phoenix 135 elpenor, Deilephila 25, 26, 27 emarginata, Sphinx 220 Embothrium 84 Emex australis 138, 378 Emex spinosa 138, 378 emilia, Angonyx 69 emu bush 98, 154, 156 Encyrtidae 30, 31, 166, 176, 193, 208, 210, 269, 374, 375 endoparasitoids 30, 32 ensifolia, Curculigo 132, 377 entomopathogenic fungi 28, 56, 138 Enyo anceus 49 Enyo cinnamomea 52, 53 Enyo lugubris 25 epicranium (of larva) 14, 16 Epilobium 260, 378 billardierianum 147, 378 glabellum 147 epiphysis 7, 8 Epipremnum 118 pinnatum 120, 376 equestris, Choerocampa 254 equestris, Sphinx 238, 254 equestris, Theretra 254 eras, Chaerocampa 125 eras, Chromis erotus 125 eras, Darapsa 125 eras, Deilephila 125 eras, Gnathothlibus 9, 27, 31, 32, 36, 37, 44, 45, 122, 123, 124, 125–128, 373, 376, 377, 379, 380, 381 eras, Gnathothlibus erotus 125 eras, Gnathothlibus erotus var. 125 Eremophila 96, 156, 233, 235 bowmanii 98, 379 deserti 98, 379 freelingii 98, 379 latrobei 98, 379 longifolia 98, 154, 156, 379 mitchellii 98, 235, 379 platycalyx 90, 380 rubra var. exotrachys 90 saligna 98, 380 sturtii 98, 380 willsii 90, 380 eremophilae, Coenotes 25, 27, 36, 40, 42, 94, 95–98, 159, 372, 376, 377, 378, 379, 380 eremophilae, Phlegethontius 95 eremophilae, Sphinx 93, 95, 96 ericifolia, Banksia 106, 378 29/08/19 11:12:19.49 INDEX erotoides, Gnathothlibus 121, 125 erotus, Gnathothlibus 122, 125 erotus cramptoni, Chromis 125 erotus eras, Chromis 125 erotus eras, Gnathothlibus 125 erotus var. eras, Chaerocampa 125 errans, Macroglossa 176 errans, Macroglossum 3, 29, 35, 36, 37, 41, 43, 166, 169, 171, 176–179, 183, 185, 194, 197, 374, 378, 379 errans f. interrupta, Macroglossum hirundo 176 errans, Macroglossum hirundo 176 erubescens, Convolvulus 64, 377 Ervatamia angustisepala 115 Ervatamia orientalis 75 Erythraeidae 29 Erythrophleum chlorostachys 84 Escallonia rubra var. macrantha 128, 377 Escalloniaceae 122, 128, 377 esculenta, Colocasia 138, 260, 268, 376 eucalyptophylla, Parsonsia 236 Eucalyptus 84 elata 102, 377 saligna 102, 377 tereticornis 102, 377 eucoxa 7 eugeniae, Josephinia 98, 378 Eulophidae 30, 31, 52, 151, 244, 372, 373, 375 Eumorpha, anchemolus 122 Eumorpha, fasciatus 25 Eumorpha, typhon 29 EUPANACRA 7, 118, 122 regularis 118 splendens 118, 120 splendens burica 119 splendens makira 118 splendens novobritannica 119 splendens paradoxa 119 splendens salomonis 118 splendens splendens 36, 37, 43, 118–120, 373, 376 splendens vellalavella 118 Eupelmidae 30, 31, 107, 115, 125, 372, 373, 374 Eupelminae 31 Euphorbia 25 Euphorbiaceae 156, 277 euphorbiae, Hyles 23, 25 euphorbiae, Sphinx 156 Euplectromorpha 31, 244, 375 Euplectrus 31, 151, 373 bicolor 31 euroa, Cypa decolor 35, 36, 108–109 europaea, Olea 219, 378 EURYPTERYX 121 bhaga 121 molucca 36, 121 molucca niepelti 121 molucca obiana 121 Exallage auricularia 132 excellens darlingtoni, Panacra 211 excellens, Angonyx 211 excellens, Panacra 210, 211 excellens, Pseudoangonyx 34, 36, 70, 210, 211 excelsa, Dendrocnide 241, 263, 380 excelsa, Lissopimpla 32, 107, 372 excelsior, Fraxinus 219, 378 exigua, Psilogramma 35, 36, 39, 41, 166, 211, 212, 213, 217, 220–223, 377, 378 Exorista 33 flaviceps 98 norrisi 135, 373 190613 Hawkmoths of Australia 3pp.indd 405 psychidivora 159, 372, 373 Exoristinae 33 Exoristini 33 exserta, Borreria 145, 378 eyes 6, 8 Fabaceae 58, 94, 98, 159, 277, 377 Fagaceae 65, 108, 109, 277 Fagraea 232, 377 berteroana 232, 377 cambagei 232, 377 false sandalwood 98 farnesiana, Acacia 98 fasciatum, Rhamphoschisma 166 fasciatus, Eumorpha 25 fecundity 33, 76, 159 feeding (adult) 6, 26 feeding (larva) 24, 28, 29 female accessory glands 13 female reproductive system 12–14 femur 7, 8, 16 ferruginea, Cypa 108 firmata, Chaerocampa 257 firmata, Theretra 257 firmata, Theretra oldenlandiae 3, 257 fitzalanii, Atractocarpus 79, 81, 378 fitzalanii, Psychotria 128, 379 fitzalanii, Randia 81 flagelliforme, Typhonium 266, 376 flagellomere 6 flagellum 6, 8 flesh flies (Sarcophagidae) 128, 373 floribundum, Clerodendrum 61, 98, 163, 165, 219, 223, 225, 232, 377 floridensis, Amphion 25 Florina 258, 263 oldenlandiae 257 silhetensis [intersecta] 263 foodplants (overview) 25 Forcipomyia 30 forewing(s) 7, 9, 17 fragans, Dracaena 120, 148, 169, 184, 204 fragans, Osmanthus 219, 378 Fraxinus angustifolia 219, 378 excelsior 219, 378 griﬃthii 219, 378 oxycarpa 219, 378 velutina 219, 378 freelingii, Eremophila 98, 379 frenulum 7, 9 friction scales 7, 10 frons 6, 8, 16 frontoclypeus 14, 16 frutescens, Ampelocissus 128, 138, 246, 260, 380 frutescens, Hodgkinsonia 169, 379 fruticosa, Polyscias 98, 376 Fuchsia (fuchsia) 27, 147, 249, 260, 378 fulvicaudata, Macroglossum corythus 170 fulviventris, Blepharipa 138, 257, 373, 375 funebris f., Hippotion brennus 132, 133 funebris, Hippotion brennus 132 fuscata, Theretra oldenlandiae 257 genital scars 18 gerstmeieri, Psilogramma 220 gigantea, Daphnis 110 gigantea, Deilephila 110 gloriosa, Daphnis 110 gloriosa, Psilogramma 212, 226 gnathos 10, 11 GNATHOTHLIBUS 7, 13, 14, 15, 118, 121–122 405 australiensis 5, 35, 36, 37, 44, 45, 122–125, 126, 127, 128, 373, 377 eras 9, 27, 31, 32, 36, 37, 44, 45, 122, 123, 124, 125–128, 373, 376, 377, 379, 380, 381 erotoides 121, 125 erotus 122, 125 erotus eras 125 heliodes 122 godarti, Agrius 24, 26, 36, 39, 41, 58, 60, 62–65, 372, 376, 377 godarti, Diludia 62 godarti, Herse 62 godarti, Sphinx 62 gonopore 11, 12 Gracillariidae 29, 31 griseola, Panacra 148 gynandromorph 371 Galium 147 leiocarpum 147, 378 gangetica, Asystasia 189 Garcinia 68 oblongifolia 68 Gardenia (gardenia) 27, 72, 75, 79, 84, 172, 173 augusta 75, 79, 84 jasminoides 75, 79, 84, 379 megasperma 79, 379 ochreata 75, 84 ovularis 84, 379 gardineri, Ampelocissus 138, 260, 380 Gentianaceae 232, 377 gibbosa, Adansonia 26 gimpi gimpi 263 glabellum, Epilobium 147 glabrata, Vitex 223, 377 Gmelina dalrympleana 232, 377 godetia 260 golden dew-drop 219 golden guinea tree 128 golden guinea vine 128 golden pothos 120 gramminifolium, Stylidium 138 grandiflora, Campsis 219, 376 grandis, Pisonia 151, 378 granitica, Pavetta 75, 379 grape 27, 28, 126, 128, 138, 241 edible 55, 128, 159 Ganzin Glory 55, 128 heart-leaved 55, 128, 249 ivy 249 ornamental 55, 58, 260 wild 52, 55, 58, 128, 249, 260, 266 wine 55, 58, 128, 159, 249, 260 Grevillea caleyi 106, 378 ‘Ivanhoe’ 106, 378 longifolia 106, 378 robusta 106, 378 griﬃthii, Fraxinus 219, 378 Guettarda speciosa 75, 79, 112, 172, 173, 275, 379 gum forest red 102 Queensland blue 102 river peppermint 102 Sydney blue gum 102 Sydney red gum 102 Gynochthodes jasminoides 93, 145, 178, 192, 193, 249, 379 Gyrocarpus 216 americanus 96, 98, 213, 216 hackeri, Carcelia 374 hainana, Acosmeryx anceus 49 hainanensis, Amplypterus panopus 68 29/08/19 11:12:19.61 40 6 HAWKMOTHS OF AUSTRALIA hainanensis, Psilogramma 220 hairy psychotria 178, 196 Hakea 84, 106, 378 halmaherana, Acosmeryx anceus 49 Haltichellinae 31 happy plant 120, 184, 204 harpe 10, 11 Hathia, lucasii 250 Hathia, tenebrosa 250 hayati, Psilogramma discistriga 220 hearing 6–7 hederifolia, Ipomoea 61, 377 Hedyotis 132, 145, 147, 260, 266, 379 heliodes, Gnathothlibus 122 heliophila, Macroglossum 35, 193 heliophila heliophila, Macroglossum 193 heliophila queenslandi, Macroglossum 193 heliophila, Macroglossum divergens 193 Hemarina 34 Hemaris 7, 11, 15, 72 bucklandii 82 cunninghami 76 janus 79 kingii 82 Hernandiaceae 98, 216, 377 herrichii, Theretra 266 Herse 58 convolvuli 59 convolvuli ab. extincta 59 convolvuli aksuensis 59 convolvuli f. posticoconflua 59 convolvuli marshallensis 59 convolvuli peitaihoensis 59 godarti 62 hespera, Nephele 36, 39, 43, 205–208, 209, 210, 374, 376 hespera, Sphinx 205 hespera f. didyma, Nephele 206 hespera f. hespera, Nephele 206 hespera var. morpheus, Nephele 205 hesperus, Choerocampa 112 hesperus, Daphnis 112 Heteropoda 193 Hexamermis cavicola 29 Hibbertia scandens 128, 135, 138, 260, 377 Hibiscus panduriformis 98, 377 hidalguense, Citharexylum 219, 380 hindwing(s) 7, 9, 17 HIPPOTION 7, 11, 128–129, 131, 132, 133, 135, 139, 143, 146, 149, 238 beddoesii 148 boerhaviae 37, 38, 44, 128, 129– 132, 142, 143, 373, 377 brennus 3, 35, 37, 38, 44, 128, 129, 132–135, 138, 139, 189, 373, 377, 378, 379 brennus f. brennus 132 brennus f. funebris 132, 133 brennus f. johanna 138 brennus f. rubribrenna 132, 133 brennus f. viettei 133 brennus funebris 132 brennus johanna 138 brennus viettei 132 celerio 4, 23, 27, 28, 33, 37, 44, 128, 129, 135–138, 159, 160, 245, 373, 376, 377, 378, 379, 380, 381 celerio ab. brunnea 135 celerio ab. pallida 135 celerio ab. unicolor 135 celerio f. luecki 136 celerio f. rosea 136 celerio f. sieberti 135 depictum 129, 141, 142 insignis 129, 238 japenum 149 190613 Hawkmoths of Australia 3pp.indd 406 johanna 3, 11, 26, 35, 37, 39, 128, 129, 133, 138–141, 373, 378 noel 148 novaebrittaniae 132 obanawae 148 ocys 135 queenslandi 260 rosetta 37, 39, 44, 128, 129, 130, 131, 132, 135, 141–145, 189, 252, 373, 377, 378, 379 rubribrenna 132 scrofa 4, 24, 26, 27, 37, 38, 44, 45, 128, 129, 145–148, 376, 378, 379, 380 taiwanensis 149 turneri 269 velox 25, 26, 32, 37, 44, 128, 129, 148–151, 373, 378 velox ab. tainanensis 149 velox tainanensis 149 hirundo, Macroglossum 3, 35, 176, 177, 197 hirundo errans f. interrupta, Macroglossum 176 hirundo errans, Macroglossum 176 hirundo f. interrupta, Macroglossum 176 hirundo hirundo, Macroglossum 176, 196 hirundo lifuensis, Macroglossum 176, 196 hirundo tonganum, Macroglossum 196 hirundo vitiense, Macroglossum 176, 196 Hodgkinsonia 169 frutescens 169, 379 hoﬀmanni, Megacorma 204 honeysuckle 128 cape 219 Japanese 219 HOPLIOCNEMA 3, 6, 7, 14, 22, 24, 25, 26, 34, 88, 89, 90, 151–152, 233 brachycera 15, 36, 37, 43, 89, 90, 91, 151, 152–154, 155, 156, 373, 379 lacunosa 5, 24, 35, 36, 40, 43, 89, 151, 152, 153, 154–156 melanoleuca 151, 152 ochra 88, 151 horsfieldii, Daphnis 112 humile, Jasminum 219, 378 hylas, Cephonodes 7, 23, 24, 25, 27, 31, 35, 71, 72, 73, 76, 77 hylas, Sphinx 71 hylas australis, Cephonodes 3, 72 hylas cunninghami, Cephonodes 76 hylas hylas, Cephonodes 72 hylas melanogaster, Cephonodes 72 hylas virescens, Cephonodes 72 HYLES 7, 23, 156–157 biguttata 157 euphorbiae 23, 25 lineata 157, 160 lineata livornicoides 157 livornica 23, 157 livornicoides 24, 25, 26, 27, 37, 45, 98, 156, 157–160, 373, 376, 377, 378, 380, 381 Hymenocallis 62 littoralis 26 hyperparasitoids 30 hypoglauca, Cissus 128, 380 hypothous, Daphnis 110 hypothous crameri, Daphnis 35, 36, 109, 275 hypothous hypothous, Daphnis 275 hypothous moorei, Daphnis 110 hypothous pallescens, Daphnis 110 Hypoxidaceae 132, 377 hyssopifolia, Ludwigia 249, 260, 266, 378 ‘Ivanhoe’, Grevillea 106, 378 Ichneumonidae 31, 32, 107, 204, 372, 374 Ichneumonoidea 30, 31, 32 idiobiont parasitoids 30 ignea, Chaerocampa 145 ignea, Theretra 145 IMBER 5, 34, 160 tropicus 5, 34, 36, 39, 43, 160–163, 374, 380 Impatiens 138, 147, 249, 251, 260, 376 balsamina 138, 147, 249, 260 oliveri 138, 147, 249, 260 walleriana 138, 147, 249, 260 inconspicua, Macroglossa 190 inconspicuoides, Sturmia 193, 251 increta f. eburnea, Psilogramma menephron 220 increta, Psilogramma 27, 212 indica, Ipomoea 61, 65, 138, 377 indica, Lagerstroemia 249, 251, 377 indica, Leea 128, 243, 249, 380 indica, Mangifera 68 indica, Protoparce convolvuli 59 indicum, Oroxylum 205 indicum, Sesamum 98, 378 indistincta bismarcki, Theretra 242 indistincta, Choerocampa 241 indistincta, Oreus 241 indistincta, Theretra 238, 241, 242 indistincta indistincta, Theretra 3, 37, 38, 44, 238, 239, 240, 241–244, 375, 380 indistincta manuselensis, Theretra 241 indistincta papuensis, Theretra 241 inflata, Manettia 147 inophyllum, Calophyllum 68 inornata, Chaerocampa 244 inornata, Theretra 37, 44, 136, 238, 244–246, 248, 267, 380 inquilinus, Phalaena 135 insignis, Hippotion 129, 238 insignis, Theretra 129, 238 insipida papuanum, Macroglossum 186, 187 insipida, Macroglossum 5, 187 insularis, Chaerocampa 246 insularis, Oreus 246 insularis, Theretra 129, 238, 246, 247 insularis ambrymenis, Theretra 246, 247 insularis insularis, Theretra 37, 238, 246–247, 251 insularis lenis, Theretra 246 insularis mollis, Theretra 246 insularis rhesus, Theretra 246 insularis valens, Theretra 246 integrifolia, Banksia 106, 378 integrifolia, Macadamia 106, 378 intersecta, Chaerocampa 263 intersecta, Theretra 263 intersecta, Theretra pinastrina 263 intersecta, Theretra silhetensis 26, 37, 38, 44, 238, 252, 258, 263–266, 375, 376, 378, 379, 380 inusitata, Macroglossa 190 inusitata, Macroglossa prometheus 190 iorioi, Megacorma 204 Ipomoea 65 aquatica 61, 377 batatas 61, 65, 128, 138, 148, 260, 377 cairica 61, 377 hederifolia 61, 377 indica 61, 65, 138, 377 nil 61, 377 Iriperenye 160 isabeliana, Ambulyx dohertyi 66 Ixora 26, 65, 189, 275 29/08/19 11:12:19.74 INDEX coccinea 115 jacaranda 26, 62 jalapa, Mirabilis 159, 378 jamdenae, Daphnis 113 jamdenae, Deilephila 113 janus austrosundanus, Cephonodes 79 janus simplex, Cephonodes 79 janus, Cephonodes 22, 36, 40, 71, 73, 75, 77, 78, 79–82, 83, 84, 372, 378, 379 janus, Hemaris 79 japenum, Hippotion 149 japonica, Lonicera 219, 377 jasmine 27, 219 jasminoides, Gardenia 75, 79, 84, 379 jasminoides, Gynochthodes 93, 145, 178, 192, 193, 249, 379 jasminoides, Morinda 93, 145 jasminoides, Pandorea 219, 376 Jasminum didymum 223, 378 humile 219, 378 nudiflorum 219, 378 polyanthum 219, 378 volubile 219, 378 joannisi, Macroglossum 26, 36, 41, 43, 166, 167, 171, 179–181, 377 johanna f., Hippotion brennus 138 johanna, Chaerocampa 138 johanna, Hippotion 3, 11, 26, 35, 37, 39, 128, 129, 133, 138–141, 373, 378 johanna, Hippotion brennus 138 johanna, Miavia 138 johanna, Panacra 138 johanna, Theretra 138 johnstonii, Cyrtosperma 268, 376 Josephinia eugeniae 98, 378 Juglandaceae 65, 277 juxta 11, 12 kang kong 61 kangaroo vine 55, 128, 249 karnatakaensis, Amplypterus panopus 68 Kigelia africana africana 87, 376 killing jars 19 kilneri, Neisosperma 115, 376 kingi, Cephonodes 82 kingii, Cephonodes 16, 24, 27, 32, 36, 40, 62, 71, 73, 75, 77, 78, 80, 81, 82–84, 201, 274, 372, 378, 379 kingii, Hemaris 82 kingii, Macroglossum 82 kleineri, Psilogramma 220 knotweed 141, 147 koalae, Psilogramma 212, 229 koinobiont parasitoids 30 koschevnikovi, Apis 46 kuvanae, Ooencyrtus 31 labial palps 6, 7, 8, 14, 29 labrum 14, 16 lachesis, Acherontia 23, 27, 36, 39, 45, 46, 47–48 lachesis, Sphinx 47 lachesis ab. atra, Manduca 47 lachesis ab. radiata, Acherontia 47 lachesis diehli, Acherontia 47 lachesis f. fuscapex, Acherontia 47 lachesis f. pallida, Acherontia 47 lachesis f. submarginata, Acherontia 47 lachesis lachesis, Acherontia 47 Lactuca sativa 138 lacunosa, Hopliocnema 5, 24, 35, 36, 40, 43, 89, 151, 152, 153, 154–156 Lagerstroemia indica 249, 251, 377 lamella antevaginalis 13, 14 190613 Hawkmoths of Australia 3pp.indd 407 lamella postvaginalis 13, 14 Lamiaceae 48, 61, 94, 98, 163, 165, 212, 219, 223, 225, 232, 377 lanceolata, Carissa 98, 210, 376 lanceolata, Dipterocarpus 109 lanceolata, Pentas 75, 79, 128, 132, 135, 138, 141, 145, 147, 189, 249, 260, 266, 379 lanceolata, Persoonia 106, 378 lanceolatum, Santalum 98, 379 Langia 34 tropicus 5, 160 Lantana (lantana) 19, 26, 62, 65, 82, 84, 93, 115, 128, 138, 148, 151, 169, 172, 179, 184, 186, 189, 219, 229 camara 75, 79, 115, 179, 189 laotensis, Marumba timora 277 Laothoe populi 25 largest hawkmoth 86, 99, 104 Larsenaikia ochreata 75, 84, 379 larvae as pests 27 larval behaviour 24–25 larval colour morphs 25 larviparous 33 lateral adfrontal suture 14, 16 latifolia, Spermacoce 135, 145, 189, 249, 379 latreillei, Theretra 247 latreillei distincta, Theretra 250 latreillei lucasi f. distincta, Theretra 250 latreillei lucasi f. montana, Theretra 250 latreillei lucasi, Theretra 250 latreillei montana, Theretra 250 latreillei, latreillei, Theretra 247 latreillii, Chaerocampa 247 latreillii, Diludia 247 latreillii, Oreus 247 latreillii, Sphinx 247 latreillii, Theretra 3, 9, 17, 27, 30, 37, 44, 238, 244, 247–250, 267, 375, 376, 377, 378, 379, 380, 381 latreillii latreillii, Theretra 247 latreillii prattorum, Theretra 3, 250 latrobei, Eremophila 98, 379 Lauraceae 277 laxiflora, Carissa 208, 210, 376 lecourti, Daphnis protrudens 116 Lecythidaceae 238, 377 Leea 251 indica 128, 243, 249, 380 legs 7, 8 Leguminosae 47 leichhardtii, Duboisia 98, 380 leiocarpum, Galium 147, 378 lenis, Theretra insularis 246, 247 lethe, Sphinx 47 lettuce 138 LEUCOMONIA 163 bethia 5, 36, 39, 41, 163–166, 374, 377 levis, Persoonia 106, 378 lewini, Sphinx 257 lewini, Theretra oldenlandiae 3, 257, 258 lifuensis, Macroglossum hirundo 176, 196 lifuensis, Panacra 148 lifuensis, Theretra 239 light trapping 19 ligustri, Sphinx 4, 24, 25, 58 Ligustrum lucidum 219, 378 sinense 219, 378 undulatum 219, 378 lilac 219 Lilliaceae 156 lily 26 arum 136, 138, 268, 376 fire 260, 376 407 peace 120, 376 spider 26 limon, Citrus 84 lineata, Hyles 157, 160 lineata, Macroglossa 190 lineata, Macroglossum prometheus 5, 22, 27, 31, 36, 40, 43, 166, 185, 186, 190–193, 195, 374, 379 lineata livornicoides, Celerio 157 lineata livornicoides, Hyles 157 lineatum, Macroglossum prometheus 190 Lissopimpla excelsa 32, 107, 372 littoralis, Hymenocallis 26 livornica var. australasiae, Phryxus 157 livornica var. livornicoides, Phryxus 157 livornica, Deilephila 157 livornica, Hyles 23, 157 livornicoides, Celerio lineata 157 livornicoides, Deilephila 157 livornicoides, Hyles 24, 25, 26, 27, 37, 45, 98, 156, 157–160, 373, 376, 377, 378, 380, 381 livornicoides, Hyles (Danneria) 157 livornicoides, Hyles lineata 157 livornicoides, Phryxus livornica var. 157 loeﬄeri, Cerberonoton 85 Loganiaceae 25, 37, 69, 71, 167, 184, 203, 377 lollybush 163, 165 longifolia, Eremophila 98, 154, 156, 379 longifolia, Grevillea 106, 378 longifolia, Notelaea 219, 378 Lonicera 128 japonica 219, 377 loniceroides, Psychotria 128, 135, 178, 196, 249, 379 looking-glass plant 93 Lophostethus dumolinii 27 lucasi f. distincta, Theretra latreillei 250 lucasi f. montana, Theretra latreillei 250 lucasi, Theretra latreillei 250 lucasii, Chaerocampa 250 lucasii, Hathia 250 lucasii, Theretra 3, 35, 37, 238, 247, 248, 250–251 lucasii, Theretra latreillii 247, 250 lucerne 159 lucida, Coprosma 93, 378 lucida, Strychnos 203, 377 lucidum, Ligustrum 219, 378 Ludwigia 26, 251 hyssopifolia 249, 260, 266, 378 octovalvis 249, 260, 266, 378 peploides 249, 266, 378 lugubris, Enyo 25 luteata, Macroglossum 170, 172 luteotincta, Chaerocampa 239 Lythraceae 238, 249, 251, 377 Macadamia 27 integrifolia 106, 378 Macroglossa 166 alcedo 167 approximans 169 approximata 201 belinda 176 corythus 172 cunninghami 76, 79 errans 176 inconspicua 190 inusitata 190 luteata 172 lineata 190 micacea 182 milvus 166 nycteris 166 proxima 172 29/08/19 11:12:19.86 40 8 HAWKMOTHS OF AUSTRALIA pseudogyrans 201 similis 201 splendens 199 tenebrosa 199 vacillans 201 yunx 76 Macroglossina 10, 11, 12, 14, 15, 25, 29, 34, 36 Macroglossini 34, 36, 69, 72, 118 MACROGLOSSUM 4, 10, 11, 15, 19, 22, 24, 26, 37, 109, 166–167, 169, 170, 171, 174, 175, 176, 181, 183, 193, 200, 205 alcedo 11, 24, 36, 41, 43, 166, 167–169, 187, 188, 374, 379 arcuatum 172 backi 201 corythus (corythus complex) 169, 170, 171 corythus corythus 36, 166, 170 172–173, 379 corythus approximans 3, 25, 35, 36, 40, 43, 166, 169–172, 173, 179, 183, 374, 378, 379 corythus fulvicaudata 170 corythus pylene 3, 169, 170 divergens 3 divergens heliophila 193 divergens queenslandi 193 doddi 173 dohertyi 167, 175 dohertyi dohertyi 173, 174 dohertyi doddi 25, 26, 36, 39, 43, 44, 92, 166, 169, 173–176, 374, 379 dohertyi melanura 173 errans 3, 29, 35, 36, 37, 41, 43, 166, 169, 171, 176–179, 183, 185, 194, 197, 374, 378, 379 heliophila 35, 193 heliophila heliophila 193 heliophila queenslandi 193 hirundo 3, 35, 176, 177, 197 hirundo errans 176 hirundo errans f. interrupta 176 hirundo f. interrupta 176 hirundo hirundo 176, 196 hirundo lifuensis 176, 196 hirundo tonganum 196 hirundo vitiense 176, 196 insipida 5, 187 insipida papuanum 186, 187 joannisi 26, 36, 41, 43, 166, 167, 171, 179–181, 377 kingii 82 luteata 170, 172 maculatum 193 melas 194 melas melas 3, 35, 36, 166, 181–182 melas moriolum 182 melas pullius 3, 183 melas queenslandi 193 micacea 167, 182, 184 micacea micacea 25, 36, 37, 40, 41, 43, 166, 167, 171, 182–185, 200, 203, 374, 377 micacea albibase 182, 183 nox 182 nubilum 36, 39, 43, 166, 167, 185–186, 190, 192, 195, 374, 378, 379 papuanum 3, 5, 35, 36, 39, 43, 135, 166, 167, 168, 186–190, 203, 374, 379 passalus 172 prometheus 193 prometheus prometheus 36, 166, 190, 193 190613 Hawkmoths of Australia 3pp.indd 408 prometheus inusitata 190 prometheus lineata 5, 22, 27, 31, 36, 40, 43, 166, 185, 186, 190–193, 195, 374, 379 prometheus lineatum 190 queenslandi 3, 35, 36, 41, 43, 44, 166, 167, 171, 177, 181, 183, 186, 192, 193–196, 379 rectans 36, 41, 43, 166, 169, 176, 177, 194, 196–199, 379 splendens 199 stellatarum 23, 26, 27, 167 stenoxanthum 3, 35, 169, 170 tenebrosa 36, 40, 43, 166, 199–201, 378 tenebrosum 199 troglodytus 3, 35, 187 troglodytus papuanum 186, 187 ungues cheni 276 ungues 275 ungues ungues 35, 36, 166, 275–276 vacillans 25, 26, 31, 36, 37, 41, 43, 166, 167, 189, 201–204, 374, 377 macromera, Diludia 220 Macropoliana 212 macrorrhizos, Alocasia 138, 257, 260, 268, 376 Macrosila bethia 163 casuarinae 216 convolvuli 59 darius 219 discistriga 219, 220 edwardsi 235 obliqua 204 severina 85 Macrostomion 32, 52, 372 maculatum, Macroglossum 193 maculiventris, Panacra 138 magnifica, Daphnis 110 majus, Antirrhinum 219, 378 makira, Eupanacra splendens 118 malabathricum, Melastoma 125, 128, 377 male accessory glands 12 male reproductive system 12 Malvaceae 26, 61, 68, 94, 98, 277, 377 mandibles (larva) 14, 16 Manduca 7 lachesis ab. atra 47 sexta 14, 23, 24, 45 Manettia inflata 147 Manettia paraguariensis 147, 379 Mangifera caesia 68 Mangifera indica 68 mango 68 mango hawkmoth 67 manica 11, 12 manuselensis, Theretra clotho 3, 241 manuselensis, Theretra indistincta 241 margarita, Chaerocampa 251 margarita, Theretra 37, 38, 45, 238, 251–254, 258, 264, 267, 378 marginata, Banksia 106, 378 marginata, Chaerocampa 251 maritima, Cayratia 249, 380 marmorata ab. dumigani, Synoecha 233 marmorata, Phlegethontius 233 marmorata, Sphinx 233 marmorata, Synoecha 25, 36, 40, 43, 94, 233–235, 379 marshallensis, Herse convolvuli 59 MARUMBA 276–277, 278 dyras tenimberi 277 quercus 277 timora 35, 36, 277–278 timora timora 277 timora laotensis 277 mastrigti aruensis, Psilogramma 229 mastrigti, Psilogramma 229 maxillary palps 6, 14, 16 maxmouldsi, Psilogramma 22, 35, 36, 39, 41, 211, 212, 213, 223–226, 377 media, Persoonia 106, 107, 378 medial adfrontal suture 14 median vein 7, 9 medicieloi, Psilogramma 220 medicine bush 172, 178 MEGACORMA 10, 14, 58, 204 hoﬀmanni 204 iorioi 204 obliqua obliqua 36, 204–205 remota 204 schroederi 204 Meganoton 34, 85, 164, 212 megasperma, Gardenia 79, 379 Melaleuca 175 melanogaster, Cephonodes hylas 72 melanoleuca, Hopliocnema 151, 152 melanomera, Diludia 220 melanura, Macroglossum dohertyi 173 melas, Macroglossum 181, 194 melas melas, Macroglossum 3, 35, 36, 166, 181–182 melas moriolum, Macroglossum 182 melas pullius, Macroglossum 3, 181, 183 melas queenslandi, Macroglossum 193 Melastoma 123 cyanoides 125, 377 malabathricum 125, 128, 377 Melastomataceae 122, 125, 128, 377 Melia 251 Meliacae 251 mellifera, Apis 27, 46, 47 Memecylaceae 167, 181 Memecylon 181 pauciflorum 181, 377 menephron, Psilogramma 2, 12, 13, 35, 212, 216, 226, 228, 375 menephron, Sphinx 211 menephron increta f. eburnea, Psilogramma 220 menephron menephron f. fasciata, Psilogramma 229 menephron menephron f. obscura, Psilogramma 226 menephron nebulosa, Psilogramma 8, 12, 13, 30, 36, 39, 41, 211, 212, 213, 220, 226–229, 375, 377, 378 Mermithidae 29 meron 7, 8 Merremia dissecta 61, 377 mesoscutellum 7, 8 mesoscutum 7, 8 mesosternum 16, 17 mesothorax 7, 14 Metamimas 98 australasiae 98, 99 banksiae 99 metapyrrha f., Nephele subvaria 208 metapyrrha, Nephele 208 metapyrrha, Zonilia 208 metascutellum 7, 8 metascutum 7, 8 metathoracic plate 17 metathorax 7, 14, 17 Metopsilus procne 250 Mexican clover 145, 260 Miavia, johanna 138 micacea, Macroglossa 182 micacea, Macroglossum 167, 182, 184 micacea albibase, Macroglossum 182, 183 micacea micacea, Macroglossum 25, 36, 37, 40, 41, 43, 166, 167, 171, 182–185, 200, 203, 374, 377 29/08/19 11:12:19.99 INDEX Microgasterinae 32 Microplitis 32, 203, 216, 375 Microplitis basalis 84, 260, 372 micropyle 14, 15 midges (Ceratopogonidae) 30 midtarsal comb 8, 10 migration 26 mindanaoensis, Amplypterus panopus 68 ming aralia 98 minimus, Protoparce 95, 96 minor, Strychnos 71, 377 mira, Panacra 269 Mirabilis jalapa 159, 378 mirabilis, Nepenthes 145, 377 mirror plant 93 miskini, Acosmeryx 27, 36, 40, 44, 48, 56–58, 372, 380, 381 miskini, Daphnusa 56 mitchellii, Eremophila 98, 235, 379 mites 27, 29–30 mixtura, Acosmeryx 49 mixtura, Zonilia 49 mollis, Theretra insularis 246, 247 molucca niepelti, Eurypteryx 121 molucca obiana, Eurypteryx 121 molucca, Eurypteryx 36, 121 molucca, Pachylia 121 Monstera deliciosa 120, 376 montana f., Theretra latreillei lucasi 250 montana, Theretra latreillei 250 montanum, Myoporum 98 moorei, Daphnis 36, 37, 43, 45, 109, 110–112, 113, 116, 275, 372, 378, 379 moorei, Daphnis hypothous 110 moorei, Darapsa 4, 110 Moraceae 205, 277 morganii praedicta, Xanthopan 26 Morinda 91, 93 citrifolia 22, 128, 167, 172, 173, 178, 186, 192, 193, 201, 379 jasminoides 93, 145 salmonensis 93, 201 moriolum, Macroglossum melas 182 morning glory 61, 65 moroides, Dendrocnide 263, 380 morpheus var., Nephele hespera 205 morpheus, Nephele 205 morpheus, Sphinx 205 morpheus, Zonilia 205 morphs (in larva) 25 morta, Acherontia atropos 47 moulting 24 mouthparts (adult) 6, 8 mouthparts (larva) 14, 16 muelleriana, Alstonia 112, 115, 376 mulla mulla 95, 98 multiple parasitoidism 30 muricata, Annona 87 muricolor, Theretra 261 Mussaenda 98, 379 myoporoides, Duboisia 98, 380 Myoporum montanum 98 Myrmecodia 173, 175, 176 beccarii 93, 175, 379 platytyrea antoinii 128, 175, 379 tuberosa 175, 379 Myrtaceae 26, 99, 102, 115, 128, 184, 186, 250, 377 nebulosa, Psilogramma menephron 8, 12, 13, 30, 36, 39, 41, 211, 212, 213, 220, 226–229, 375, 377, 378 Neisosperma kilneri 115, 376 nematodes 29 Nemoraea 33, 257, 375 Nemoraeini 33 Neogurelca 11, 69 Nepenthaceae 129, 145, 377 Nepenthes mirabilis 145, 377 rowanae 145, 377 tenax 145, 377 NEPHELE 109, 167, 205, 206 accentifera 205 chiron 205 didyma 205 didyma ab. hespera 205 didyma f. didyma 206 didyma f. hespera 205 hespera 36, 39, 43, 205–208, 209, 210, 374, 376 hespera var. morpheus 205 metapyrrha 208 morpheus 205 subvaria 31, 36, 39, 43, 205, 206, 207, 208–210, 374, 376 subvaria f. metapyrrha 208 subvaria f. subvaria 208 Nephelium 251 neriastri, Choerocampa 115 nerii, Daphnis 23, 30, 109 nerii, Sphinx 109 Nerium 275 nessus, Chaerocampa 254 nessus, Pergesa 254 nessus, Sphinx 238, 254 nessus, Theretra 26, 27, 29, 238, 254, 255 nessus albata, Theretra 254 nessus nessus, Theretra 37, 44, 238, 254–257, 375, 376, 377 nessus var. rubicundus, Chaerocampa 254 nestor, Meganoton, 204 nestor, Sphinx, 204 Netelia 32, 107, 372 Newcastelia spodiotricha 98, 377 niepelti, Eurypteryx molucca 121 nil, Ipomoea 61, 377 nitens, Tetrastigma 128, 138, 249, 251, 380 Noctuidonema 29 noel, Hippotion 148 norrisi, Exorista 135, 373 Notelaea longifolia 219, 378 Notonagemia 85 novaebrittaniae, Hippotion 132 novobritannia, Ambulyx dohertyi 65, 66 novobritannica, Eupanacra splendens 119 novoirlandensis, Ambulyx dohertyi 66 nox, Macroglossum 182 Ntyarlke 160 nubilum, Macroglossum 36, 39, 43, 166, 167, 185–186, 190, 192, 195, 374, 378, 379 nuclear polyhedrosis virus (NPV) 28 nudiflorum, Jasminum 219, 378 Nyctaginaceae 129, 138, 151, 156, 159, 238, 254, 377 nycteris, Macroglossa 166 naga, Acosmeryx 49 native mulberry 240, 263 Nauclea orientalis 79, 112, 379 Neanastatinae 31 nebulosa, Diludia 226 nebulosa, Meganoton 226 nebulosa, Psilogramma 226 obanawae, Hippotion (Chaerocampa) 148 obiana, Eurypteryx molucca 121 obliqua, Diludia 204 obliqua, Macrosila 204 obliqua, Meganoton 204 obliqua obliqua, Megacorma 36, 204–205 190613 Hawkmoths of Australia 3pp.indd 409 409 obliterans, Perigonia 205 oblonga, Cissus 55, 58, 128, 147, 241, 246, 249, 260, 380 oblongifolia, Garcinia 68 obovatus, Ptilotus 95, 98, 376 ocellata, Smerinthus 25 ocelli 6, 14 ochra, Chelacnema 3, 5, 8, 24, 25, 34, 36, 40, 43, 88–91, 151, 152, 154, 155, 379, 380 ochra, Hopliocnema 35, 88, 151 ochreata, Gardenia 75, 84 ochreata, Larsenaikia 75, 84, 379 Ochrosia elliptica 115, 376 octopunctata, Sphinx 129 octovalvis, Ludwigia 249, 260, 266, 378 ocys, Hippotion 135 odorata, Cananga 87, 376 odorata, Canthium 75, 79, 81, 84, 178 odorata, Psydrax 75, 79, 81, 84, 178, 379 Oecophylla smaragdina 29 Oldenlandia auricularia 132 oldenlandiae, Chaerocampa 257 oldenlandiae, Deilephila 257 oldenlandiae, Florina 257 oldenlandiae, Sphinx 257 oldenlandiae, Theretra 4, 15, 23, 27, 32, 37, 39, 44, 238, 250, 252, 257–260, 264, 267, 269, 270, 271, 375, 376, 377, 378, 379, 380, 381 oldenlandiae, Xylophanes 257 oldenlandiae firmata, Theretra 3, 257 oldenlandiae fuscata, Theretra 257 oldenlandiae lewini, Theretra 3, 257 oldenlandiae oldenlandiae, Theretra 3, 257, 258 oldenlandiae olivascens, Theretra 257 oldenlandiae samoana, Theretra 3, 257 Olea africana 219, 378 europaea 219, 378 paniculata 219, 378 Oleaceae 212, 219, 223, 228, 378 oleifolia, Psydrax 84, 379 oleifolium, Canthium 84 olivacens, Theretra 257 olivascens, Theretra oldenlandiae 257 olive common 219 fragrant 219 northern 228 oliveri, Impatiens 138, 147, 249, 260 ommatidia 6 Onagraceae 129, 147, 156, 238, 249, 251, 260, 266, 378 oocytes 13 Ooencyrtus 31, 166, 176, 193, 208, 210, 269, 374, 375 kuvanae 31 opaca, Clematicissus 128, 138, 246, 249, 260, 380 Oreocallis 84 Oreus 238, 241, 242, 246, 247, 266 Oreus 238, 241, 242, 246, 247, 266 indistincta 241 insularis 246 latreillii 247 tryoni 266 orientalis, Ervatamia 75 orientalis, Nauclea 79, 112, 379 orientalis, Protoparce 59 orientalis, Tabernaemontana 75, 115, 376 Oroxylum indicum 205 Osmanthus fragans 219, 378 ostium bursae 12, 13 Otopheidomenidae 29, 30 outer margin 7, 9 29/08/19 11:12:20.11 410 HAWKMOTHS OF AUSTRALIA ovaries 13 ovarioles 13 ovata, Carissa 210, 376 ovipore 13 ovolarviparous 33 ovularis, Gardenia 84, 379 Oxyambulux dohertyi queenslandi 65 wildei 66 oxycarpa, Fraxinus 219, 378 oxycarpum, Abutilon 61 Paederia 276 paeoniifolius, Amorphophallus 260, 376 pagoda flower 232 Palexorista 33, 98, 138, 141, 145, 159, 179, 186, 189, 372, 373, 374 pallescens, Daphnis 110 pallescens, Daphnis hypothous 110 pallescens, Deilephila 110 pallicosta, Choerocampa 4 pallida ab., Hippotion celerio 135 pallida f., Acherontia lachesis 47 pallida, Chaerocampa 244 pallida, Theretra 244 Panacra dohertyi 118 excellens 210, 211 excellens darlingtoni 211 griseola 148 johanna 138 lifuensis 148 lignaria 148 maculiventris 138 mira 269 pseudovigil 148 rosea 148 splendens 118 splendens splendens 118 turneri 269 pandacaqui, Tabernaemontana 115, 376 pandorana, Pandorea 219, 376 Pandorea jasminoides 219, 376 Pandorea pandorana 219, 376 panduriformis, Hibiscus 98, 377 paniculata, Olea 219, 378 paniculatum var. syncarpum, Coelospermum 93, 186, 201, 378 paniculatum, Clerodendrum 232, 377 panopus, Calymnia 68 panopus, Compsogene 68 panopus, Compsogene (Calymnia) 68 panopus, Sphinx 67, 68 panopus celebensis, Amplypterus 68 panopus hainanensis, Amplypterus 68 panopus karnatakaensis, Amplypterus 68 panopus mindanaoensis, Amplypterus 68 panopus panopus, Amplypterus 35, 36, 67–69 panopus panopus, Compsogene 68 panopus seramensis, Amplypterus 68 panopus sumbawanensis, Amplypterus 68 papaya, Carica 26, 62, 79, 138, 151, 169, 179, 184, 189, 210, 260, 266 papillae anales 13 papuana, Angonyx 69, 377 papuana, Angonyx testacea 69 papuana bismarcki, Angonyx 69 papuana papuana f. serrata, Angonyx 69 papuana papuana, Angonyx 36, 40, 45, 69–71, 211 papuanum, Macroglossum 3, 5, 35, 36, 39, 43, 135, 166, 167, 168, 186–190, 374, 379 papuanum, Macroglossum insipida 187 papuanum, Macroglossum troglodytus 187 190613 Hawkmoths of Australia 3pp.indd 410 papuensis, Psilogramma 35, 36, 39, 41, 211, 212, 213, 227, 229–232, 376, 377 papuensis, Theretra 241 papuensis, Theretra clotho 3, 241 papuensis, Theretra indistincta 241 paradoxa, Eupanacra splendens 119 paradoxus, Brachychiton 162, 380 paraguariensis, Manettia 147, 379 parasites 29 parasitoids/parasitoidism 27, 29, 30 Parsonsia 235, 236, 237 eucalyptophylla 236 plaesiophylla 237 sankowskyana 237 straminea 237, 376 Parthenocissus 251 quinquefolia 52, 128, 138, 249, 380 tricuspidata 128, 138, 249, 380 passalus, Macroglossum 172 Passifloraceae 58 patagia 7, 8 patatas, Sphinx 59 pathogens 28 pauciflorum, Memecylon 181, 377 paukstadtorum, Psilogramma 220 Pavetta 275 australiensis 75, 79, 178, 379 brownii 75, 379 granitica 75, 379 pavonica, Calymnia 68 pavonicus, Amplypterus 68 pawpaw 26, 62, 138, 151, 169, 179, 184, 189, 210, 260, 266 pearsalli, Anastatus 31 Pedaliaceae 94, 98, 378 pedicel 6, 8 Pediobius, atamiensis 31 Pediobius, bruchicida 31 pedunculatum, Crinum 26 peitaihoensis, Herse convolvuli 59 penninervis, Cissus 128, 380 Pentas (pentas) 19, 26, 27, 128, 141, 145, 169, 178, 189, 249 lanceolata 75, 79, 128, 132, 135, 138, 141, 145, 147, 189, 249, 260, 266, 379 penumbra, Psilogramma 35, 36, 211, 212, 213, 232–233 peploides, Ludwigia 249, 266, 378 pepper vine 249 perfumed canthium 75, 79, 81, 84, 178 Pergesa 15, 238 nessus 254 vampyrus 129 Perigonia obliterans 205 Perigonia testacea 69 Persicaria decipiens 26, 141, 378 Persicaria prostrata 147, 378 Persoonia lanceolata 106, 378 levis 106, 378 media 106, 107, 378 pest (adult and larva) 27 Phalaena inquilinus 135 phallus 11, 12 phasianinus, Centropus 219 Philampelini 34, 72 Philidris cordata 175 philippinensis, Acosmeryx anceus 49 philippinensis, Cerberonoton rubescens 86 philodice, Colias 25 Phlegethontius convolvuli 59 distincta 62 eremophilae 95 marmorata 233 phoenix, Chaerocampa 251 phoenix, Elpenor 135 phoenyx, Sphinx 148 Phoridae 33 photinophylla, Dendrocnide 241, 263, 380 Phryxus livornica var. australasiae 157 Phryxus livornica var. livornicoides 157 picus, Cephonodes 3, 35, 71, 72, 73, 76, 77 pilifer 6, 8 pinastrina intersecta, Theretra 263 pinnatum, Epipremnum 120, 376 Pipturus argenteus 240, 241, 263, 380 Pisonia 149, 151 aculeata 151, 378 grandis 151, 378 umbellifera 151, 378 placida, Daphnis 112 placida, Darapsa 112 placida placida, Daphnis 36, 40, 43, 45, 109, 111, 112–115, 116, 372, 376, 377 placida placida, Deilephila 113 placida salomonis, Daphnis 113 plaesiophylla, Parsonsia 237 Planchonia careya 260, 377 planta 15 Plantaginaceae 212, 219, 378 platycalyx, Eremophila 90, 380 Platygastroidea 30, 31, 32 platytyrea antoinii, Myrmecodia 128, 175, 379 pleuron 8, 10 plowmanii, Anthurium 120, 376 Plumeria acutifolia 138, 376 Plumeria rubra 138, 376 pluto, Xylophanes 30 Podranea brycei 138, 376 Podranea ricasoliana 138, 219, 376 poliostemma, Psychotria 178, 199, 379 pollination 26 polyanthum, Jasminum 219, 378 Polygonaceae 58, 129, 138, 141, 147, 156, 238, 249, 378 Polymeria 61, 377 Polyscias fruticosa 98, 376 populi, Laothoe 25 porcellus, Deilephila 4 porcellus, Sphinx 4 porcia, Deilephila 145 Portulaca 159, 378 Portulacaceae 58, 156, 159, 378 posterior apophysis 13 posterior cubital vein 7, 9 potentia, Chaerocampa 260 praedicta, Xanthopan morganii 26 Prasadiseius 29 prattorum, Theretra 250 prattorum, Theretra latreillii 3, 247, 250 predators 28–29 pretarsal claws 8, 10, 16 pretarsus 10 prickly saltwort 64 primary parasitoids 30 privet 27 box-leaf 219, 378 broad-leaf 219, 378 common 219, 378 privet hawkmoth 4, 216 proboscis (adult) 6, 8, 26 proboscis (pupa) 15, 17, 26 procne, Chaerocampa 250 procne, Choerocampa 250 procne, Metopsilus 250 procne, Theretra 250 prolegs 15 prometheus, Macroglossum 193 29/08/19 11:12:20.24 INDEX prometheus inusitata, Macroglossum 190 prometheus lineata, Macroglossum 5, 22, 27, 31, 36, 40, 43, 166, 185, 186, 190–193, 195, 374, 379 prometheus lineatum, Macroglossum 190 prometheus prometheus, Macroglossum 36, 166, 190, 193 prominens, Carcelia 159, 373 Prostanthera 98, 377 althoferi 98, 377 striatiflora 98, 377 prosternum 16, 17 prostrata, Persicaria 147, 378 Proteaceae 99, 106, 107, 378 prothoracic shield 15, 16 prothoracic spiracle 17 prothorax 7, 14, 15 Protoparce convolvuli 59 convolvuli ab. fasciata 59 convolvuli indica 59 distans 59 minimus 95 orientalis 59 protrudens, Choerocampa 115 protrudens, Daphnis 111, 113, 115, 116, 379 protrudens, Deilephila 115 protrudens lecourti, Daphnis 116 protrudens protrudens, Daphnis 36, 37, 40, 43, 45, 109, 115–118, 372 proxima, Deilephila 257 proxima, Macroglossa 172 proxima, Theretra 257 PSEUDOANGONYX 5, 10, 34, 210-211 excellens 34, 36, 70, 210, 211 pseudoconvolvuli, Sphinx 59 pseudogyrans, Macroglossa 201 pseudovigil, Panacra 148 PSILOGRAMMA 5, 10, 11, 15, 24, 25, 26, 34, 35, 88, 164, 211–213, 214, 217, 219, 220, 221, 224, 226, 227, 228, 230, 232, 235, 374 anne 212, 226 argos 11, 32, 35, 36, 39, 41, 211, 212, 213–216, 375, 377 casuarinae 17, 25, 27, 35, 36, 39, 41, 166, 211, 212, 213, 216–219, 221, 226, 229, 375, 376, 377, 378, 380 choui 220 danneri 220 discistriga discistriga 35, 36, 211, 212, 219–220, 227 discistriga hayati 220 exigua 35, 36, 39, 41, 166, 211, 212, 213, 217, 220–223, 377, 378 gerstmeieri 220 gloriosa 212, 226 hainanensis 220 hausmanni 212, 216 increta 27, 212 kleineri 220 koalae 212, 229 mastrigti 229 mastrigti aruensis 229 maxmouldsi 22, 35, 36, 39, 41, 211, 212, 213, 223–226, 377 medicieloi 220 menephron 12, 13, 35, 212, 216, 226, 228, 375 menephron increta f. eburnea 220 menephron menephron f. fasciata 229 menephron nebulosa 8, 12, 13, 30, 36, 39, 41, 211, 212, 213, 220, 226–229, 375, 377, 378 190613 Hawkmoths of Australia 3pp.indd 411 nebulosa 226 papuensis 35, 36, 39, 41, 211, 212, 213, 227, 229–232, 376, 377 paukstadtorum 220 penumbra 35, 36, 211, 212, 213, 232–233 stameri 220 stameri choui 220 surholti 220 psilosperma, Strychnos 184, 377 Psithyros 166 psychidivora, Exorista 159, 372, 373 Psychotria 75, 169, 177, 194, 196, 197, 379 daphnoides 178, 379 fitzalanii 128, 379 loniceroides 128, 135, 178, 196, 249, 379 poliostemma 178, 199, 379 Psydrax 184 attenuata 84, 273, 379 odorata 75, 79, 81, 84, 178, 379 oleifolia 84, 379 ridigula 273, 379 Ptilotus obovatus 95, 98, 376 pubescens, Boerhavia 159, 254, 378 puellaris, Chaerocampa 257 puellaris, Theretra 257 pullius, Macroglossum melas 3, 181, 182, 183 pulvilli 8, 10 puncture vine 159 pylene, Macroglossum corythus 3, 169, 170 Pyralidae 31 quadrifida, Coprosma 93, 378 quaterna, Sphinx 205 queenslandi, Ambulyx dohertyi 36, 65–66, 67 queenslandi, Chaerocampa 260 queenslandi, Hippotion 260 queenslandi, Macroglossum 3, 35, 36, 41, 43, 44, 166, 167, 171, 177, 181, 183, 186, 192, 193–196, 379 queenslandi, Macroglossum divergens 193 queenslandi, Macroglossum heliophila 193 queenslandi, Macroglossum melas 193 queenslandi, Oxyambulyx dohertyi 65 queenslandi, Theretra 15, 25, 37, 44, 238, 239, 240, 242, 247, 260–263, 375, 380 Quercus 109 quercus, Marumba 277 quinquefolia, Parthenocissus 52, 128, 138, 249, 380 racemosa, Aidia 79, 81, 378 Radermachera sinica 219, 377 radial sector vein 7, 9 radicans, Campsis 219, 376 radiosa, Theretra 260, 261 radius 7, 9 ramiflorus, Chionanthus 228, 378 Randia fitzalanii 79, 81, 378 Randia sessilis 81, 378 Raphidophora australasica 120, 376 ratstail 98 rearing larvae 22–23 rectans, Macroglossum 36, 41, 43, 166, 169, 176, 177, 194, 196–199, 379 regularis, Eupanacra 118 remota, Megacorma obliqua 204 Remusatia vivipara 268, 376 reniformis, Cissus 243, 246, 380 repens, Cissus 128, 241, 249, 380 repens, Coprosma 93, 147, 178, 378 411 repens, Duranta 82, 84, 219, 380 reticulatum, Coelospermum 135, 172, 178, 179, 260, 378 retinaculum 7, 9 rhababarum, Rheum 138, 378 Rhagastis 238 Rhamnaceae 277 Rhamphoschisma 166 fasciatum 166 scottiarum 176 rhesus, Theretra 250 rhesus insularis, Theretra 246 Rheum rhababarum 138, 378 rhombifolia, Cissus 128, 249, 380 Rhopalopsyche 166 rhubarb 27, 138 ricasoliana, Podranea 138, 219, 376 Richardia brasiliensis 145, 260, 379 Richardia scabra 145, 260, 379 ridigula, Canthium 273 ridigula, Psydrax 273, 379 riparia, Calliandra 189 river peppermint 102 robusta, Grevillea 106, 378 Rogadinae 32 roly-poly 64, 159 rosacea, Daphnis torenia 112 Rosaceae 212, 219, 277 rosea f., Hippotion celerio 136 rosea, Panacra 148 roseafasciata, Sphinx convolvuli 59 rosetta, Chaerocampa 141 rosetta, Hippotion 37, 39, 44, 128, 129, 130, 131, 132, 135, 141–145, 189, 252, 373, 377, 378, 379 rowanae, Nepenthes 145, 377 rubescens, Cerberonoton 3, 35, 85 rubescens, Diludia 84 rubescens, Meganoton 85 rubescens philippinensis, Cerberonoton 86 rubescens rubescens, Meganoton 86 rubescens severina, Cerberonoton 34, 85 rubescens severina, Meganoton 85 rubescens thielei, Cerberonoton 86 rubescens titan, Cerberonoton 86 Rubia tinctorum 147, 379 Rubiaceae 25, 37, 48, 72, 75, 79, 81, 84, 91, 93, 94, 98, 109, 112, 117, 118, 122, 128, 129, 132, 135, 138, 141, 145, 147, 156, 167, 169, 172, 173, 175, 178, 184, 186, 189, 192, 193, 196, 199, 201, 205, 238, 249, 260, 266, 271, 273, 275, 276, 378 rubra, Plumeria 138, 376 rubra var. exotrachys, Eremophila 90, 380 rubra var. macrantha, Escallonia 128, 377 rubribrenna f., Hippotion brennus 132 rubribrenna, Hippotion 132 rufescens severina, Meganoton 85 Rumex 138, 378 acetosa 249, 378 Rutaceae 251 Sabiaceae 277 sacculus 10, 11 saccus 10, 11 Salicaceae 156, 277 salicifolia, Veronica 219 378 saligna, Eremophila 98, 380 saligna, Eucalyptus 102, 377 salmonensis, Morinda 93, 201 salomonis, Daphnis placida 113 salomonis, Eupanacra splendens 118 Salsola 64, 159, 376 samoana, Theretra oldenlandiae 3, 257, 258 29/08/19 11:12:20.36 412 HAWKMOTHS OF AUSTRALIA sankowskyana, Parsonsia 237 Santalaceae 94, 98, 379 Santalum acuminatum 98, 379 Santalum lanceolatum 98, 379 Sapindaceae 251, 277 sapor, Chaerocampa 125 Sarcophagidae 33, 128, 373 Sarcorohdendorfia alcicomis 128, 373 satanas, Acherontia 47 sativa, Lactuca 138 sativa, Medicago 159 Saurauia andreana 128, 376 sausage tree 88 scabra, Richardia 145, 260, 379 scandens, Hibbertia 128, 135, 138, 260, 377 scape 6, 8 Scelionidae 31, 32, 112, 115, 118, 125, 128, 151, 172, 179, 193, 372, 373, 374 schauﬀelbergeri, Ambulyx 65 schlechtendalii, Anthurium 120, 376 scholaris, Alstonia 112, 115, 121, 376 schroederi, Megacorma 204 scottiarum, Rhamphoschisma 176 scrofa, Chaerocampa 145 scrofa, Deilephila 145 scrofa, Hippotion 4, 24, 26, 27, 37, 38, 44, 45, 128, 129, 145–148, 376, 378, 379, 380 scrofa, Theretra 145 Scrophulariaceae 88, 90, 94, 98, 152, 154, 156, 233, 235, 379 sculpta, Cizara 91 secondary parasitoids 30 seminal vesicle 12 Senometopia 269, 375 sepium, Calystegia 61 seramensis, Amplypterus panopus 68 serrata, Banksia 106, 378 serrulata, Candollea 138 sesame 98 Sesamum indicum 98, 378 Sesia cunninghami 76 sesquipedale, Angraecum 26 sessilis, Atractocarpus 81, 378 sessilis, Randia 81 severina, Cerberonoton 3, 24, 34, 35, 36, 39, 41, 58, 84, 85–88, 104, 376, 377 severina, Cerberonoton rubescens 34, 85 severina, Macrosila 85 severina, Meganoton 85 severina, Meganoton rubescens 85 severina, Psilogramma 85 sexta, Manduca 14, 23, 24 sexual communication 26 Sichiini 36, 277 sight 26 signum 13 silhetensis [intersecta], Florina 263 silhetensis intersecta, Theretra 26, 37, 38, 44, 238, 252, 258, 263–266, 375, 376, 378, 379, 380 silhetensis silhetensis, Theretra 264, 266 silhetensis, Theretra 15, 32, 252, 263, 267 silkpod 237 silver bush 95, 98 silver-striped hawkmoth 135 similis, Macroglossa 201 simplex, Cephonodes janus 79 sinense, Ligustrum 219, 378 sinica, Radermachera 219, 377 sinuatus, Stenocarpus 106, 378 Siphonini 33 smaragdina, Oecophylla 22 Smerinthinae 10, 11, 12, 34, 36 190613 Hawkmoths of Australia 3pp.indd 412 Smerinthini 6, 7, 14, 27, 34, 36 Smerinthus dyras 276 Smerinthus ocellata 25 Smicromorphinae 31 smooth-barked apple 102 snake vine 128, 135, 260 snakeweed 26, 98, 151, 179, 184, 189, 193 snapdragon 219 sobria, Chaerocampa 257 sojejimae, Acherontia 47 Solanaceae 47, 48, 58, 94, 98, 380 solomonis, Ambulyx dohertyi 65, 66 soursop 87 Sparassidae 193, 269 Spathiphyllum wallisii 120, 376 Spathodea campanulata 88, 205, 219, 228, 229, 232, 377 speciosa, Guettarda 75, 79, 112, 172, 173, 379 spectabilis, Bougainvillea 151, 378 Spectrum charon 47 Spermacoce 189, 275 latifolia 135, 145, 189, 249, 379 spermatheca 13 spermathecal gland 13 Sphecodina abbottii 25 Sphinginae 10, 11, 12, 14, 15, 25, 34, 36 Sphingini 7, 10, 34, 36 Sphingonaepiopsis 11, 69 Sphingulini 6, 14, 34, 36 Sphinx 4, 14, 25, 45, 58 abadonna 59 anceus 48, 49 ardenia 91 ardeniae 91 argentata 257 atropos 45 australasiae 98, 99 boerhaviae 129 brennus 132 castaneus 103 casuarinae 216 celerio 128, 135 chiron 205 cingulata 58 convolvuli 58, 59 convolvuli ab. alicea 59 convolvuli roseafasciata 59 convolvuli var. batatae 59 convolvuli var. distans 59 convolvuli var. nigricans 59 didyma 205 distincta 62 drancus 257 emarginata 220 equestris 238, 254 eremophilae 93, 95, 96 euphorbiae 156 godarti 62 hespera 205 hylas 71 lachesis 47 latreillii 247 lethe 47 lewini 257 ligustri 4, 24, 25 marmorata 233 menephron 211 morpheus 205 nerii 109 nessus 238, 254 nestor 204 octopunctata 129 oldenlandiae 257 panopus 67, 68 patatas 59 phoenyx 148 porcellus 4 pseudoconvolvuli 59 quaterna 205 stellatarum 166 substrigilis 65 tisiphone 135 triangularis 98, 103 vampyrus 129 velox 148 vigil 148 spilota, Deilephila 250 spinarum, Carissa 208 spines 10 spinnerets 14, 16 spinosa, Emex 138, 378 spinosum, Xanthium 147 spinulosa, Banksia 106, 378 spiracles (adult) 7, 8, 10 spiracles (larva) 14, 16 spiracles (pupa) 17, 18 spiracular furrows 17, 18 splendens, Angonyx 118 splendens, Eupanacra 118, 119, 120 splendens, Macroglossa 199 splendens, Macroglossum 199 splendens burica, Eupanacra 119 splendens makira, Eupanacra 118 splendens novobritannica, Eupanacra 119 splendens paradoxa, Eupanacra 119 splendens salomonis, Eupanacra 118 splendens splendens, Eupanacra 36, 37, 43, 118–120, 373, 376 splendens splendens, Panacra 118 splendens vellalavella, Eupanacra 118 spodiotricha, Newcastelia 98, 377 spurs 8, 10 square stem 135, 145, 189, 249 Stachytarpheta, cayennensis 26, 98, 151, 179, 184, 189, 193, 380 stameri chuai, Psilogramma 220 stameri, Psilogramma 220 stans, Tecoma 219, 377 star-cluster 128 stellatarum, Macroglossum 23, 26, 27, 167 stellatarum, Sphinx 166 stemmata 14, 16 Stenocarpus sinuatus 106, 378 stenoxanthum, Macroglossum 3, 35, 169 Sterculiaceae 160, 162, 380 sterigma 13, 14 Stictocardia tiliifolia 61, 377 stinging tree, giant 263 stinging tree, shiny-leafed 263 stinkwood 98, 216 straminea, Parsonsia 237, 376 striatiflora, Prostanthera 98, 377 strychnine tree 203 Strychnos 25, 37, 69, 182, 184 lucida 203, 377 minor 71, 377 psilosperma 184, 377 Sturmia 33 convergens 62, 372 inconspicuoides 193, 251 Sturmiini 33 sturtii, Eremophila 98, 380 Stylidium gramminifolium 138 styx, Acherontia 27, 46, 47 subcostal vein 7, 9 subdentata, Acosmery anceus 49 subvaria, Nephele 31, 36, 39, 43, 205, 206, 207, 208–210, 374, 376 subvaria f. metapyrrha, Nephele 208 subvaria f. subvaria, Nephele 208 29/08/19 11:12:20.48 INDEX subvaria, Zonilia 208 sumatrana, Winthemia 93, 115, 128, 210, 372, 373, 374 sumbawanensis, Amplypterus panopus 68 super parasitoidism 32 surholti, Psilogramma 220 swamp mahogany 128 sweet potato 61, 65, 128, 148, 260 swinhoei f., Hippotion velox 149 swinhoei, Chaerocampa 148 SYNOECHA 14, 34, 151, 233 marmorata 25, 36, 40, 43, 94, 233–235, 379 marmorata ab. dumigani 233 Syringa vulgaris 219, 378 Syzygium tierneyanum 26, 115, 128, 184, 186, 250 Tabernaemontana 275 divaricata 115, 376 orientalis 75, 115, 376 pandacaqui 115, 376 Tachinidae 33 Tachininae 33 tainanensis ab., Hippotion velox 149 tainanensis, Hippotion velox 149 taishanensis, Cordyceps 28 taiwanensis, Hippotion 149 tar vine 159 Tarenna 84, 379 dallachiana 79, 379 taro 260, 268 tarsomeres 8, 10 tarsus 7, 8, 10 Tecoma stans 219, 377 Tecomaria capensis 219, 377 tegulae 7 tegumen 10, 11 Telenominae 32 Telenomus 32, 112, 115, 118, 125, 128, 151, 172, 179, 193, 372, 373, 374 remus 33 Temnora 109, 167, 205 tenax, Nepenthes 145, 377 tenebrosa, Chaerocampa 250 tenebrosa, Hathia 250 tenebrosa, Macroglossa 199 tenebrosa, Macroglossum 36, 40, 43, 166, 199–201, 378 tenebrosa, Theretra 250 tenebrosum, Macroglossum 199 tenimberi, Marumba dyras 277 tereticornis, Eucalyptus 102, 377 terrestris, Tribulus 159, 381 tersa, Xylophanes 25 testacea, Angonyx 69 testacea, Perigonia 69 testacea papuana, Angonyx 69 testes 12 TETRACHROA 10, 34, 164, 235 edwardsi 11, 36, 39, 45, 235–238, 376 tetraphylla, Deplanchea 232, 376 Tetrastichinae 52, 372 Tetrastigma nitens 128, 138, 249, 251, 380 thelyotokous parthenogenesis 30 THERETRA 7, 12, 15, 129, 238, 242, 246, 247, 250, 258, 260, 261, 263, 266, 269 alecto 23, 238 amara 247 aquila 263 brennus 132 capensis 238 celata 35, 239 190613 Hawkmoths of Australia 3pp.indd 413 celata celata 37, 38, 44, 238, 239–241, 242, 380 celata babarensis 239 celerio 135 cleopatra 241 cloacina 239 clotho 239, 240, 241 clotho celata 239 clotho manuselensis 3, 241 clotho papuensis 3, 241 curvilinea 241 deserta 247 equestris 254 firmata 257 herrichii 266 ignea 145 indistincta 238, 241 indistincta indistincta 3, 37, 38, 44, 238, 239, 240, 241–244, 375, 380 indistincta bismarcki 242 indistincta manuselensis 241 indistincta papuensis 241 inornata 37, 44, 136, 238, 244–246, 248, 267, 380 insignis 129, 238 insularis 129, 238, 246, 247 insularis insularis 37, 238, 246–247, 251 insularis ambrymenis 246, 247 insularis lenis 246, 247 insularis mollis 246, 247 insularis valens 246, 247 intersecta 263 johanna 138 latreillei 247 latreillei distincta 250 latreillei lucasi 250 latreillei lucasi f. distincta 250 latreillei lucasi f. montana 250 latreillei montana 250 latreillii 3, 9, 17, 27, 30, 37, 44, 238, 244, 247–250, 267, 375, 376, 377, 378, 379, 380, 381 latreillii lucasii 247, 250 latreillii prattorum 3, 247, 250 lifuensis 239 lucasii 3, 35, 37, 238, 247, 248, 250–251 margarita 37, 38, 45, 238, 251–254, 258, 264, 267, 378 muricolor 261 nessus 26, 27, 29, 238, 254, 255 nessus albata 254, 255 nessus nessus 37, 44, 238, 254–257, 375, 376, 377 oldenlandiae 4, 15, 23, 27, 32, 37, 39, 44, 238, 250, 252, 257–260, 264, 267, 269, 270, 271, 375, 376, 377, 378, 379, 380, 381 oldenlandiae firmata 3, 257 oldenlandiae fuscata 257 oldenlandiae lewini 3, 257, 258 oldenlandiae oldenlandiae 3, 257, 258 oldenlandiae olivascens 257 oldenlandiae samoana 3, 257, 258 olivacens 257 pallida 244 papuensis 241 pinastrina intersecta 263 prattorum 250 procne 250 proxima 257 puellaris 257 queenslandi 15, 25, 37, 44, 238, 239, 240, 242, 247, 260–263, 375, 380 radiosa 260, 261 413 rhesus 250 rhesus insularis 246 scrofa 245 silhetensis 15, 32, 238, 252, 263, 267 silhetensis silhetensis 264, 266 silhetensis intersecta 26, 37, 38, 44, 238, 252, 258, 263–266, 375, 376, 378, 379, 380 tenebrosa 250 tryoni 37, 44, 238, 244, 245, 248, 250, 266–269, 375, 376 turneri 37, 39, 44, 129, 238, 250, 255, 258, 267, 269–271, 375, 377, 380 velox 148 walduckii 247 thielei, Cerberonoton rubescens 85, 86 thorax 7, 8, 14 thuringiensis, Bacillus (BT) 28 tibia 7, 8, 10 Tibouchina urvilleana 125, 377 tierneyanum, Syzygium 26, 115, 128, 184, 186, 250 tiliifolia, Stictocardia 61, 377 timon, Timonius 117, 379 Timonius timon 117, 379 timora, Marumba 35, 36, 277–278 timora laotensis, Marumba 277 timora timora, Marumba 276, 277 Timoria 58 tinctorum, Rubia 147, 379 tisiphone, Sphinx 135 titan, Cerberonoton rubescens 86 tomentosum, Clerodendrum 26, 219, 377 tonganum, Macroglossum hirundo 196 torenia rosacea, Daphnis 112 tornus 7, 9 Tortricidae 31 Toxicodendron vernicifluum 68 tracyanum, Clerodendrum 232, 377 transtilla 10 transversa, Dioscorea 271, 377 triangularis, Acherontia 103 triangularis, Brachyglossa 103 triangularis, Coequosa 7, 15, 23, 24, 27, 32, 36, 37, 43, 98, 99, 102, 103–108, 372, 378 triangularis, Sphinx 98, 103 Tribulus terrestris 159, 381 Trichogramma 32, 154, 169, 184, 204, 210, 250, 271, 372, 373, 374 carverae 151, 373, 375 pretiosum 32, 151, 373 Trichogrammatidae 30, 31–32 tricuspidata, Parthenocissus 128, 138, 249, 380 trifolia, Cayratia 128, 260, 271, 380 trifolia, Vitex 219, 223, 377 trigger-plant 138 trochanter 7, 8, 16 troglodytus papuanum, Macroglossum 186 troglodytus troglodytus, Macroglossum 187 troglodytus, Macroglossum 3, 35, 187 tropicus, Imber 5, 34, 36, 39, 43, 160–163, 374, 380 tropicus, Langia 5, 160 true legs 15, 16 trumpet vine, Argentine 219, 260 trumpet vine, Chinese 219 tryoni, Chaerocampa 266 tryoni, Oreus 266 tryoni, Theretra 37, 44, 238, 244, 245, 248, 250, 266–269, 375, 376 tuberculatus, Dipterocarpus 109 tuberosa, Myrmecodia 175, 379 tumidity 15, 16 29/08/19 11:12:20.61 414 HAWKMOTHS OF AUSTRALIA turneri, Hippotion 269 turneri, Panacra 269 turneri, Theretra 37, 39, 44, 129, 238, 250, 255, 258, 267, 269–271, 375, 377, 380 twirly-whirly tree 98, 216 typhon, Eumorpha 29 Typhonium angustilobum 260, 376 brownii 138, 376 flagelliforme 266, 376 umbellifera, Pisonia 151, 378 Uncaria 275 uncus 10, 11 undulatum, Ligustrum 219, 378 ungues cheni, Macroglossum 276 ungues ungues, Macroglossum 35, 36, 166, 275–276 unguiculata, Clarkia 260, 378 unicolor, Cephonodes 79 unicolor ab., Agrius convolvuli 59 unicolor ab., Hippotion celerio 135 uniformis, Cypa 109 Urticeae 238, 241, 263, 380 urvilleana, Tibouchina 125, 377 Utnerrengatye 160 vacillans, Macroglossa 201 vacillans, Macroglossum 25, 26, 31, 36, 37, 41, 43, 166, 167, 189, 201–204, 374, 377 vacillans backi, Macroglossum 201 vacillans vacillans, Macroglossum 201 valens, Theretra insularis 246, 247 Valerianaceae 156 Vampyronema 29 vampyrus, Pergesa 129 vampyrus, Sphinx 129 variegata ab., Agrius convolvuli 59 variegatum, Meganoton 235 vas deferentia 12 vein(s) 7, 9 vellalavella, Eupanacra splendens 118 velox ab. tainanensis, Hippotion 149 velox tainanensis, Hippotion 149 velox, Chaerocampa 148 velox, Hippotion 25, 26, 32, 37, 44, 128, 129, 148–151, 373, 378 190613 Hawkmoths of Australia 3pp.indd 414 velox, Sphinx 148 velox, Theretra 148 velutina, Fraxinus 219, 378 ventral prolegs 15, 16 Verbenaceae 47, 58, 94, 98, 212, 219, 277, 380 vernicifluum, Toxicodendron 68 Veronica (veronica) 219, 378 salicifolia 219, 378 vertex 6, 8 vesica 11, 12 viettei f., Hippotion brennus 133 viettei, Hippotion brennus 132 vigil, Sphinx 148 villosum, Alangium 115, 377 vinculum 10, 11 vinifera x rupestris, Vitis 55, 58, 128, 138, 249, 260, 381 vinifera, Vitis 55, 58, 128, 138, 159, 243, 249, 260, 380 virescens, Cephonodes hylas 72 viruses 28, 30 Vitaceae 24, 25, 28, 49, 52, 55, 58, 118, 122, 128, 129, 138, 145, 147, 156, 159, 238, 240, 241, 242, 243, 246, 249, 251, 260, 266, 271, 380 Vitex acuminata 223, 377 glabrata 223, 377 trifolia 219, 223, 377 vitiense, Macroglossum hirundo 176, 196 Vitis 49, 251 vinifera 55, 58, 128, 138, 159, 243, 249, 260, 380 vinifera x rupestris 55, 58, 128, 138, 249, 260, 381 vivipara, Remusatia 268, 376 vojtechi, Zacria 5, 25, 34, 35, 36, 40, 45, 271, 272–274, 379 volubile, Jasminum 219, 378 vulgaris, Syringa 219, 378 walduckii, Chaerocampa 247 walduckii, Theretra 247 walkeri, Amphimoea 6 walleriana, Impatiens 138, 147, 249, 260 wallisii, Spathiphyllum 120, 376 water primrose 266 Wendlandia 275 white-lined hawkmoth 157 wildei, Ambulyx 36, 65, 66–67 wildei, Oxyambulyx 66 willsii, Eremophila 90, 380 wing markings 7, 9 wing veins 7, 9 Winthemia 33, 107, 163, 374 Winthemia sumatrana 93, 115, 128, 210, 372, 373 Winthemiini 33 Wolbachia 28 wonga-wonga vine 219 Xanthium spinosum 147 Xanthopan 7 Xanthopan morganii praedicta 26 xanthus, Cephonodes 72 Xylophanes 157 chiron 205 drancus 257 oldenlandiae 257 pluto 30 tersa 25 valvae 10, 11 varroa mites 27 yellow bells 219 Yeperenye 27, 160 ylang ylang tree 87 yorkii, Choerocampa 148 yunx, Macroglossa 76 ZACRIA 5, 34, 271–272 vojtechi 5, 25, 34, 35, 36, 40, 45, 271, 272–274, 379 Zantedeschia 88 aethiopica 136, 138, 260, 268, 376 Zonilia antipoda 208 ardenia 91 chiron 205 metapyrrha 208 mixtura 49 morpheus 205 subvaria 208 Zosterops lateralis 228 Zygobothria 33, 166, 229, 374, 375 atropivora 219, 375 Zygophyllaceae 156, 159, 381 29/08/19 11:12:20.70

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