Main Insect Morphology and Phylogeny: A textbook for students of entomology

Insect Morphology and Phylogeny: A textbook for students of entomology

, , ,
This textbook provides an in-depth treatment of the structures and the phylogeny of the megadiverse Hexapoda. It presents an up-to-date overview of general insect morphology with detailed drawings, scanning electron micrographs, and 3-D reconstructions and is a modern synthesis of insect systematics. The work is an invaluable reference for students and researchers of biology and is a must for evolutionary biologists.

Comprehensive treatment of insect morphology with numerous detailed illustrations, brilliant electron micrographs and 3-D reconstructions Glossary for quick reference Overview of traditional and modern techniques in insect morphology

Aims and Scope:

In the last decades a remarkable renaissance has materialized in insect morphology, mainly triggered by the development of new cutting-edge technologies. This is an exciting time for biological synthesis where the mysteries and data derived from genomes can be combined with centuries of data from morphology and development. And, now, more than ever, detailed knowledge of morphology is essential to understanding the evolution of all groups of organisms. In this “age of phylogenomics” researchers rely on morphological data to support molecular findings, test complex evolutionary scenarios, and for placing fossil taxa. This textbook provides an in-depth treatment of the structures and the phylogeny of the megadiverse Hexapoda. The first part presents an up-to-date overview of general insect morphology with detailed drawings, scanning electron micrographs, and 3-D reconstructions. Also included is a chapter covering innovative morphological techniques (e.g., µ-computer tomography, 3-D modeling), brief treatments of insect development and phylogenetic methods, and a comprehensive morphological glossary. The second part is of a modern synthesis of insect systematics that includes taxon-specific morphological information for all Orders. The work is an invaluable reference for students and researchers working in all facets of biology and is a must for evolutionary biologists. A detailed understanding of morphology is essential in unraveling phylogenetic relationships and developing complex evolutionary scenarios. Increasingly researchers in phylogenomics are re/turning to morphological data to support their findings, while the development of new cutting-edge technologies has further increased interest in this growing field. This definitive handbook provides an in-depth treatment of insect morphology. The first part presents an up-to-date overview of insect morphology with detailed drawings, brilliant scanning electron micrographs and 3-D reconstructions as interactive PDFs. Thisis complemented by a chapter on innovative morphological techniques (e.g., µ-computer tomography, 3-D modeling) and a comprehensive morphological glossary. The second part treats the state of the art in insect systematics and includes taxon-specific morphological information for all orders. Systematicsare treated formally, with for example the arguments for relationships (“apomorphies”) always listed explicitly. The work is a useful reference for students and researchers working in different fields of biology and a must for those dealing with insects from an evolutionary perspective.
Categories: Biology\\Zoology
Year: 2014
Publisher: Walter de Gruyter
Language: english
Pages: 533
ISBN 10: 3110262630
ISBN 13: 9783110262636
Series: de Gruyter Textbook
File: PDF, 58.83 MB
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Beutel, Friedrich, Ge, Yang
Insect Morphology and Phylogeny
De Gruyter Graduate

Rolf G. Beutel, Frank Friedrich, Si-Qin Ge,
Xing-Ke Yang

Insect Morphology and
Phylogeny

A textbook for students of entomology

Authors
Rolf G. Beutel
Friedrich-Schiller-Univ., Institut f.
Spezielle Zoologie u. Evolutionsbiologie
Erbertstr. 1
07743 Jena
Germany

Si-Qin Ge
Chinese Academy of Sciences
Institute of Zoology
1 Beichen West Road, Chaoyang
100101 Beijing
China

Frank Friedrich
Universität Hamburg
Biozentrum Grindel u. Zoologisches Museum
Martin-Luther-King-Platz 3
20146 Hamburg
Germany

Xing-Ke Yang
Chinese Academy of Sciences
Institute of Zoology
1 Beichen West Road, Chaoyang
100101 Beijing
China

ISBN 978-3-11-026263-6
e-ISBN 978-3-11- 026404-3
Library of Congress Cataloging-in-Publication data
A CIP catalog record for this book has been applied for at the Library of Congress.
Bibliographic information published by the Deutsche Nationalbibliothek
The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliografie;
detailed bibliographic data are available in the Internet at http://dnb.dnb.de.
© 2014 Walter de Gruyter GmbH, Berlin/Boston
Typesetting: Werksatz Schmidt & Schulz GmbH, Gräfenhainichen
Printing and binding: Hubert & Co. GmbH & Co. KG, Göttingen
Cover image: Mengenilla moldrzyki Pohl et al., 2012 (Strepsiptera, Mengenillidae)
Nikon D 90, 25 mm Zeiss Luminar macro lense, adjustable extension bellows, flashlight.
Combined stack of several partially focused images using Helicon Focus. Courtesy H. Pohl
♾ Printed on acid-free paper
Printed in Germany
www.degruyter.com

The carpet makers of Isfahan deliberately knot tiny flaws into their rugs,
because perfection is an attribute reserved for God.
We dedicate this book to Niels-Peder Kristensen who has set a shining example
in insect morphology and phylogeny.

Foreword
This book emerged from a close cooperation between scientists from the Institute of
Zoology of the Chinese Academy of Sciences (CAS) and two German institutions, the
Institut für Spezielle Zoologie und Evolutionsbiologie mit Phyletischem Museum of
the Friedrich-Schiller-Universität Jena and the Biozentrum Grindel & Zoologisches
Museum of the University of Hamburg. Between these institutions, joint research projects have focused on insect anatomy and innovative morphological techniques and
on the phylogeny and evolutionary history of different hexapod lineages. Our progress and interest in these topics are reflected in the contents of this work.
The tremendous importance of Hexapoda was highlighted in numerous contributions and will not be treated in detail here. However, the most outstanding feature
of this clade is its unparalleled diversity. With approximately 1,000,000 described
species, they comprise more than half of the known total species diversity on this
planet. However, what is presently known is apparently only the tip of the iceberg.
Estimates of the real diversity range between 2 million species and a staggering
number of 30 million. Hexapod species often occur in extremely dense local populations and can form an immense biomass. Up to 100,000 springtails in only one m3 of
forest topsoil or millions of mosquitos forming gigantic swarms are only two examples of such incredible population density, among many others. Hexapods are largely
and primarily missing in marine habitats, but they play a crucial role in nearly all
terrestrial ecosystems and occur in a broad variety of limnic habitats. They have a
huge impact on human health as vectors of many diseases (e.g., malaria, sleeping
sickness), and many species are important plant pests or pests of stored products.
Positive aspects of hexapods include their role as predators or parasitoids of pest
species (mostly pest insects) and as pollinators of plants including important crops.
Insects are an important food source for numerous animal species and traditionally
also for humans in many parts of the world. Last but not least, the production of silk
and honey have been important economic factors going back several thousand years.
The combination of unusually complex morphology, fascinating biology, remarkable
species attractiveness and charisma, economic and medical impact, and various other
aspects have made hexapods a highly attractive group for researchers and dedicated
amateurs for centuries. Moreover, the grave threat posed by an unparalleled biodiversity crisis to the seemingly inexhaustible hexapod diversity presents one more very
serious reason to intensify the study and detailed documentation of this fascinating
group of organisms.
Insect morphology was a flourishing discipline in the first two thirds of the 20th
century, with outstanding researchers such as J. Chaudonneret and H. Weber in European countries, but also excellent entomologists in other parts of the world. Morphology based systematic entomology arguably reached a peak with the publication of
Willi Hennig’s groundbreaking work “Die Stammesgeschichte der Insekten” in 1969.
Towards the end of the 20th century, the detailed anatomical study of insects became

VIII

Foreword

less and less popular, a development apparently linked to the rise of molecular systematics. However, in the last ten years innovative techniques and new theoretical
concepts (e.g., “evolutionary morphology”) have led to a remarkable renaissance of
the investigation of structures and functions of Hexapoda.
Molecular systematics has “evolved” with breathtaking momentum in the last ten
years (see e.g., 1KITE.org). Robust “molecular phylogenies” will likely be available for
Hexapoda and other groups of organisms in the very near future. Nevertheless, morphology will continue to play a vital role for different reasons. First of all, it provides
an independent source of information for critically evaluating molecular trees (and
vice versa), a procedure referred to as the “model of reciprocal enlightenment” by
W. Hennig. Organisms cope with their environment using their morphological structures, which are the main target of natural selection. Body functions cannot be understood without solid morphological data, and detailed and meaningful evolutionary
scenarios cannot be developed without knowing the changes on the phenotypic level.
Another obvious reason is that morphology is the only source of information regarding fossils. To reconstruct the evolution of Hexapoda in its historical dimension is
only possible using morphological data for the placement of extinct taxa.
The primary purpose of this book is to provide a comprehensive overview of
hexapod morphology, mainly, but not exclusively, for investigations in an evolutionary context. On one hand an overwhelming richness of available data is made easily
accessible here, including also extensive and highly valuable sources in non-anglophone languages (see below). On the other hand, extensive results of our own morphological investigations are integrated in this volume, including comprehensive tables
of muscles with recently introduced nomenclatures, high quality SEM micrographs,
and computer-based 3D reconstructions. The second main aim is to outline the state
of the art in hexapod phylogenetics. The almost unprecedented progress in hexapod
systematics in the last years, arguably comparable to Hennig’s “Stammesgeschichte
der Insekten”, provides an almost ideal background. Long disputed questions, such
as the position of Strepsiptera (“the Strepsiptera problem”), are now settled, and it is
probably not overoptimistic to assume that a more or less completely resolved hexapod
phylogeny (on the interordinal level) will be available in the very near future. In this
context it should be emphasized that this is not only owed to the immense progress
in molecular systematics, but also to several coordinated morphology-based projects,
including phylogenetic studies of Polyneoptera and Holometabola.
The first main part of this book covers general hexapod morphology (1. Morphol­
ogy) which is followed by a concise treatment of the development and immature
stages (2. Reproduction, development and immature stages) and an extensive
glossary (3. Glossary of hexapod morphology). A broad spectrum of traditional and
innovative morphological techniques is described briefly in the next chapter (4. Tra­
ditional and modern morphological techniques) followed by a brief introduction
into morphology-based phylogenetics (5. Phylogenetic reconstruction based on
morphology). The second main part (6. The hexapod orders) covers all currently

Foreword

IX

recognized hexapod orders and their systematic relationships. The main focus in the
ordinal chapters is on the morphology, but these chapters also contain shorter sections on the distribution and diversity, taxonomy, biology, reproduction, fossil record,
and economic importance of the different orders.
The information presented in this volume is based on numerous sources (see
7. Literature). Works extensively used are Snodgrass’ classical “Principles of Insect
Morphology”, the German “Handbook of Zoology” series (De Gruyter), the “Traité de
Zoologie” (edited by P.P. Grassé), some textbooks in German language (e.g., “Entomologisches Praktikum”, G. Seifert), and last but not least “Evolution of the Insects”
by D. Grimaldi and M. Engel. It should be emphasized that numerous specialists have
made valuable contributions to this volume by carefully reviewing chapters (see
Acknowledgements). Few chapters were written by invited specialists Assoc. Prof. Dr.
M. Bai (Chinese Academy of Sciences), Dr. B. Wipfler, and Dipl. Biol. K. Schneeberg
(Institut für Spezielle Zoologie und Evolutionsbiologie, University Jena).
This book addresses students of entomology, especially those interested in morphology, phylogeny and evolution, but also researches dealing with hexapod systematics or other aspects of entomology. A slightly modified Chinese version of this book
is presently in preparation. We hope that this contribution will not only promote the
study and investigation of insect morphology and evolution but also stimulate international exchange and joint research projects in systematic entomology and related
disciplines.
Rolf G. Beutel
Frank Friedrich

Si-Qin Ge
Xing-Ke Yang

Acknowledgements
To finish this book project would not have been possible without the help of many
persons. First of all we would like to acknowledge the invaluable contribution made by
Hans Pohl (FSU Jena) by providing numerous SEM micrographs and macro light images
of superb quality, including the one used for the cover photo, and also extremely helpful
comments on many chapters. Numerous drawings of outstanding quality were provided
by Wenzhu Li, which is also greatly appreciated. Additional valuable images were made
available by Romano Dallai (Universita di Siena), Ryuichiro Machida, Yuta Mashimo,
Apisit Thipaksorn (Sugadaira Montane Research Center), Reinhard Predel (Universität
zu Köln), Rico Spangenberg, Lars Möckel, Katrin Friedemann and Frank Hünefeld (FSU
Jena). The chapters of this book were carefully reviewed by colleagues, almost all of
them externally. This was crucial for the success of the project. The external reviewers include Horst Aspöck (Universität Wien) (Neuropterida), Ulrike Aspöck (Naturhistorisches Museum Wien) (Neuropterida), Matt Bertone (North Carolina State University)
(Diptera), Alexander Blanke (Universität Bonn) (Odonata), Sven Bradler (Universität
Göttingen) (Phasmatodea), Daniel Burckhardt (Coleorrhyncha, Sternorrhyncha), Fabian
Haas (Dermaptera), Bruce Heming (University of Alberta) (development, Thysanoptera),
Wieland Hertel (FSU Jena) (morphology), Frank Hünefeld (FSU Jena) (Lepidoptera),
Karl Kjer (Rutgers University) (Trichoptera), K.-D. Klass (Sen­ckenberg Naturhistorische
Sammlungen Dresden) (techniques, Dictyoptera, Mantophasmatodea), Markus Koch
(University Hamburg) (entognathous orders), Günther Köhler (FSU Jena) (Orthoptera),
Lars Krogmann (Naturkundemuseum Stuttgart) (Hymenoptera), Robert E. Lewis (Iowa
State University) (Siphonaptera), Ryuichiro Machida (University of Tsukuba) (development, Archaeognatha, Zygentoma), Rudolf Meier (National University of Singapore) (claclistics), Christine Mißbach (MPI for Chemical Ecology, Jena) (central nervous system),
Carsten Müller (Universität Greifswald) (eyes), Michael Ohl (Naturkundemuseum Berlin)
(Neuroptera), Thomas Simonsen (The Natural History Museum, London) (Lepidoptera),
Adam Ślipiński (CSIRO Canberra) (Coleoptera), Hans Strümpel (Universität Hamburg)
(Sternorrhyncha, Auchenorrhyncha), Claudia Szumik (CONICET, Argentina) (Embioptera), Gert Tröster (Universität Göttingen) (Phthiraptera), Christiane Weirauch (University of California, Riverside) (Hemiptera), Brian Wiegmann (North Carolina State University) (Diptera), Kazunori Yoshizawa (University of Hokkaido) (Embioptera, Zoraptera,
Psocoptera), and Peter Zwick (MPI of Limnology, Plön) (Plecoptera). We are very grateful
for the immense support by these highly competent reviewers, which helped greatly to
improve the quality of the book. Jasmin Tannert did the lettering of all drawings. Renate
Walter helped with the preparation of SEM samples. Janin Naumann and Rommy Petersohn provided excellent microtome sections, which were important for different parts
of the book. Kathrin Streitberger helped proofreading the manuscript. This is also gratefully acknowledged, as is the multiple support with illustrations provided by Gunnar
Brehm. Finally our great thanks go to the Chinese Academy of Sciences (Beijing) and the
Friedrich-Schiller-Universität Jena for various kinds of supports for this project.

Contents
1	Morphology — 1
1.1	Integument — 1
1.1.1	Cuticle and epidermis — 1
1.1.2	Canals and pores — 3
1.1.3	Surface structures, microtrichia and setae — 4
1.1.4	Cuticular sensilla — 5
1.1.5	Scolopidia — 7
1.1.6	Integumental gland cells — 9
1.1.7	Ecdysis — 10
1.2	Head — 12
1.2.1	Segmentation, sutures and cephalic regions — 12
1.2.2	Head capsule — 13
1.2.3	Cephalic endoskeleton — 16
1.2.4	Labrum and epipharynx — 17
1.2.5	Antennae — 19
1.2.6	Mandibles — 20
1.2.7	Maxillae — 21
1.2.8	Labium — 21
1.2.9	Hypopharynx — 22
1.2.10	Salivarium — 23
1.3	Thorax — 29
1.3.1	Segmentation and composition of segments — 29
1.3.2	Prothorax — 32
1.3.3	Pterothoracic segments — 33
1.3.4	Legs — 38
1.3.5	Attachment structures — 41
1.3.6	Wings — 45
1.4	Abdomen — 59
1.4.1	General organization — 59
1.4.2	External structures of the male and female postabdomen — 62
1.4.3	Male and female internal genital organs — 63
1.5	Nervous system — 67
1.5.1	Brain — 69
1.5.2	Suboesophageal complex — 73
1.5.3	Postcephalic ganglionic chain — 74
1.5.4	Visceral nervous system (VNS) — 74
1.6	Photoreceptor organs — 75
1.6.1	Extraocular photoreception — 75
1.6.2	Compound eyes — 76
1.6.3	Ocelli — 80

XII

Contents

1.6.4	Stemmata — 81
1.7	Tracheal system — 82
1.8	Circulatory system — 87
1.8.1	Haemocoel, diaphragmata and alary muscles — 87
1.8.2	Dorsal vessel — 88
1.8.3	Accessory pulsatile organs — 91
1.8.4	Haemolymph — 91
1.9	Digestive tract — 93
1.9.1	Preoral cavity — 93
1.9.2	Foregut — 93
1.9.3	Midgut — 95
1.9.4	Hindgut — 96
1.10	Excretory organs — 97
1.11	Endocrine organs and the hormone system — 99
1.12	Fat body — 102
2	Reproduction, development and immature stages — 104
2.1	Fertilization and egg structure — 104
2.2	Embryonic development — 106
2.2.1	Cleavage and germ band formation — 106
2.2.2	Short germ and long germ embryos — 108
2.2.3	Germ layer formation and blastokinesis — 109
2.2.4	Segmentation — 111
2.2.5	Organogenesis — 112
2.3	Postembryonic development — 113
2.4	Larval and pupal forms — 115
3	Glossary — 117
3.1
Morphology — 117
3.1.1
General terms — 117
3.1.2
Integument — 117
3.1.3
Sensilla and sensory organs — 118
3.1.4
Head capsule — 119
3.1.5
Head appendages — 122
3.1.6
Thorax — 123
3.1.7
Legs — 126
3.1.8
Wings — 127
3.1.9
Abdomen — 128
3.1.10
Male genital organs — 129
3.1.11
Female genital organs — 130
3.1.12
Nervous system and associated structures — 131
3.1.13
Photoreceptor organs — 134

Contents

3.1.14
3.1.15
3.1.16
3.1.17
3.1.18
3.1.19
3.2
3.2.1
3.2.2
3.2.3
3.2.4

Tracheal system — 135
Circulatory system — 135
Digestive tract — 137
Excretory organs — 138
Endocrine organs and the hormone system — 139
Fat Body — 139
Reproduction, development and immature stages — 140
Fertilization and egg structure — 140
Cleavage and embryonic developement — 140
Postembryonic development — 141
Larval and pupal forms — 142

4	Traditional and modern techniques in insect morphology — 143
4.1	Fixation — 143
4.2	Dissection — 144
4.3	Maceration — 145
4.4	Scanning electron microscopy (SEM) — 146
4.5	Transmission electron microscopy (TEM) — 148
4.6	Histology — 150
4.7	Serial Block-Face Scanning Electron Microscopy (SBFSEM) — 152
4.8	Focused Ion Beam (FIB) — 153
4.9	Confocal laser scanning microscopy (CLSM) — 154
4.10	Micro-computer tomography (µ-CT) — 155
4.11	Computer-based 3-dimensional reconstruction — 157
4.12	Geometric morphometrics (Ming Bai) — 158
4.12.1	Terminology and principles — 159
4.12.2	Geometric morphometrics analysis — 161
5	Phylogenetic reconstruction based on morphology — 164
5.1	Hennigian terminology and principles — 164
5.2	Cladistics — 168
5.2.1	Taxon sampling — 168
5.2.2	Selection of characters — 169
5.2.3	Character state coding and building a list of characters — 170
5.2.4	Data matrix — 171
5.2.5	Parsimony analyses — 171
5.2.6	Trees, their presentation and statistics — 172
6	The orders of Hexapoda — 174
6.1	Collembola (common name: springtails) — 178
6.2	Protura (common name: coneheads) — 184
6.3	Diplura (common name: two-pronged bristletails) — 188

XIII

XIV

Contents

6.4	Archaeognatha (common name: jumping bristletails) — 196
6.5	Zygentoma (common names: silverfish and firebrats) — 201
6.6	Ephemeroptera (common name: mayflies) — 209
6.7	Odonata (common names: damselflies, dragonflies) — 217
6.8	Plecoptera (common name: stoneflies) — 229
6.9	Dermaptera (common name: earwigs) — 236
6.10	Embioptera (common name: webspinners) — 242
6.11	Phasmatodea (common names: stick and leaf insects) — 246
6.12	Orthoptera (common names: katydids = bush-crickets, crickets, grasshoppers, locusts) — 251
6.13	Zoraptera (common names: ground lice, angel insects) — 257
6.14	Grylloblattodea (common names: ice crawlers, rock crawlers, icebugs)
(Benjamin Wipfler) — 265
6.15	Mantophasmatodea (common names: heelwalkers, gladiators)
(Benjamin Wipfler) — 272
6.16	Mantodea (common names: mantises, praying mantises)
(Benjamin Wipfler) — 277
6.17	Blattodea (common names: roaches and termites)
(Benjamin Wipfler) — 282
6.18	Psocoptera (common names: barklice, booklice, barkflies) — 296
6.19	Phthiraptera (common name: true lice) — 304
6.20	Thysanoptera (common names: thrips, fringe wings) — 313
6.21	Auchenorrhyncha (common names: Cicadas, leafhoppers, planthoppers, froghoppers or spittle bugs, treehoppers) — 320
6.22	Sternorrhyncha (common name: plantlice) — 326
6.22.1	Psyllina, Psylloidea (common names: psyllids,
jumping plantlice, lerp insects) — 328
6.22.2	Aleyrodina, Aleyrodoidea (common name: whiteflies) — 331
6.22.3	Aphidina (common name: aphids) — 334
6.22.4	Coccina (common names: scale insects, mealy bugs) — 338
6.23	Coleorrhyncha (common name: moss bugs) — 341
6.24	Heteroptera (common name: true bugs) — 347
6.25	Hymenoptera (common names: sawflies, wood wasps, bees,
wasps, ants) — 364
6.26	Neuroptera (common name: net-winged insects) — 376
6.27	Megaloptera (common names: alderflies, dobsonflies, fishflies) — 385
6.28	Raphidioptera (common names: snakeflies, camelneck flies) — 393
6.29	Coleoptera (common name: beetles) — 401
6.30	Strepsiptera (common name: twisted wing parasites)
(Hans Pohl & Rolf G. Beutel) — 415
6.31	Trichoptera (common name: caddisflies) — 423
6.32	Lepidoptera (common names: moths and butterflies) — 433

Contents

XV

6.33	Mecoptera (common names for subgroups: scorpionflies, hangingflies
etc.) — 447
6.34	Siphonaptera (common name: fleas) — 457
6.35	Diptera (common name: true flies) (Katharina Schneeberg &
Rolf G. Beutel) — 465
7	Literature — 480
7.1	Textbooks and comprehensive works — 480
7.2	Review articles — 482
7.3	Cladistic software and related studies — 483
7.4	Complete references — 483
Taxonomic Index — 508

1 Morphology
1.1 Integument
1.1.1 Cuticle and epidermis
Like other euarthropods (Chelicerata, Myriapoda, crustaceans [probably paraphyletic]) Hexapoda (=insects in the widest sense)1 are characterized by a differentiated
exoskeleton formed by the external cuticle. It is composed of sclerites, membranes2
and semimembranous areas. The cuticle is a biological composite material containing chitin, proteins, lipids and catecholamines (e.g., N-acetyl-dopamine). Catechol­
amines cross-link proteins and chitin filaments, which results in specific mechanical
properties. The exoskeleton is usually robust in most areas and results in an improved
mechanical protection of the body, but it also provides differentiated attachment areas
for a complex muscular system. It is a precondition for the formation of a complex
locomotor apparatus with true articulations and complex appendages (arthropodia),
which was a key evolutionary innovation of Euarthropoda. Protection against desiccation is another function in most terrestrial arthropods, usually linked with the presence of an external wax layer (see below).
The cuticle does not only cover the surface. Endoskeletal structures are formed as
ingrowths, referred to as apodemes if they are solid and as apophyses (or entapophyses) if they are hollow. They play an important role in most hexapods, especially
as muscle attachment areas, but also increase the mechanical stability of certain body
parts, such as the tentorium or postoccipital ridge in the head, or the furcae and
pleural ridges in the thoracic segments. Internal organs such as the tracheae and
fore- and hindgut are also covered by a very thin cuticle, the intima.
The cuticle is secreted by the single-layered epidermis, which is also referred to
as hypodermis due to its position below the cuticle (Fig. 1.1.1.1). It is mainly formed
by cubic or more or less strongly flattened cells with a basal lamina (0.2–0.5 µm),
but contains also different types of gland cells (see 1.1.6 Integumental gland cells),
cells forming setae (tormogen and trichogen cells, see 1.1.4 Cuticular sensilla),
sensorial cells and oenocytes. The basal lamina is formed by epidermal cells but also
by plasmatocytes. Its main components are collagen, glycoproteins and glycosaminoglycans (Chapman 1998). In contrast to most other cells of the hypodermis, the
oenocytes have no contact with the cuticle. They are often large (more than 100 µm

1 Insects is the commonly used name for the entire Hexapoda (see title). In the text of the book we
use consistently Hexapoda/hexapods for all insects including the entognathous orders (Collembola,
Protura, Diplura) and Insecta/insects for Ectognatha, i.e. Archaeognatha, Zygentoma and Pterygota.
2 Terms in bold face in the parts 1 and 2 (Morphology and Development) are covered in the glossary
(part 3).

2

1 Morphology

in diameter) and characterized by a large nucleus, an extensive endoplasmatic reticulum, a low number of mitochondria, and crystalline inclusions. Oenocytes synthesize
hydrocarbons that contribute to the epicuticle (Chapman 1998).

set
gld

epc

brc

exc

enc

gli
glc

hypc oen

bme toc

trc

thc

sec

Fig. 1.1.1.1: Integument. Abbr.: bme – basement membrane, brc – basal ring cell, enc –
endocuticle, epc – epicuticle, exc – exocuticle, glc – gland cell, gli – glia cell, hypc –
hypodermal cell, oen – oenocyte, set – seta, sec – sensory cell, thc – thecogen cell, toc –
tormogen cell, trc – trichogen cell. Redrawn from Seifert (1995).

The unmodified epidermal cells are held together by zonulae adherens near their
apical regions and connected by septate junctions more basally. Desmosomes,
hemidesmosomes and septate junctions also occur (Seifert 1995; Chapman 1998). The
apical membrane forms a series of short ridges or projections (resembling microvilli)
which are flattened apically. These plasma membrane plaques are the sites of the
secretion of the epicuticle and chitin fibers (Chapman 1998). All epidermal cells have
a glandular function as they secrete cuticle and also enzymes involved in its production and digestion.
The cuticle is composed of three layers, the external epicuticle, the exocuticle,
and the internal endocuticle (Fig. 1.1.1.1). The two inner layers are initially secreted as
a soft and more or less homogenous procuticle by the epidermis. It contains chains of
alpha-chitin (poly-N-acetylglucosamine: [C8H13NO5]x) connected by hydrogen bonds
as larger units with a parallel arrangement, the micelles or microfibrils (2.5–3 nm).
These are embedded in a matrix of silk-like and globular proteins. The micelles lie
parallel to each other in each plane, but the arrangement differs in successive layers
of the cuticle. A regular helicoidal arrangement in a series of lamellae is a typical
pattern.
The poly-N-acetylglucosamine molecules form the main component of the procuticle. The matrix protein of the procuticle and endocuticle is the flexible and watersoluble arthropodin. In the outer layer a hardening process takes place involving


1.1 Integument

3

dehydration and tanning mediated by phenoloxidases. This transforms the arthropodin into the rigid, brownish and water-resistant sclerotin of the exocuticle. A specialized matrix material is the rubber-like, highly elastic protein resilin. It occurs in
sockets of true hairs (setae), in wings, and in mechanically highly active areas such
as for instance the wing articulations.
The thickness of the epicuticle, which forms a multilayered external barrier,
varies strongly (ca. 30 nm in culicid larvae, maximum ca. 4,000 nm). It is always free
of chitin. In pterygote insects it is covered by a wax layer secreted by oenocytes. It is
composed of paraffins and esters, which reduce water loss via evaporation. Its thickness varies between 10 nm and 1,000 nm and different surface modifications can
occur (e.g., as whitish dust in Aleyrodoidea [white flies] and Coniopterygidae [dustywings]). An additional external cement layer occurs in some groups of insects (e.g.,
Blattaria), in some cases as an open meshwork. The very thin intermediate lamina of
the epicuticle (ca. 15 nm) is mainly formed by the hardened protein cuticulin, which
is similar to the sclerotin of the exocuticle. The homogenous inner layer is called the
dense lamina. It is highly robust mechanically.
The exocuticle is strongly developed in the sclerites of the exoskeleton and can
be half as thick as the entire cuticle in some cases. It is very strong under compressive forces, but comparatively weak under tension. It is very thin in the membranous
areas, which are mainly formed by the endocuticle, which is flexible and able to
resist tensile forces. Membranous areas occur at articulations but also on other body
regions in most groups, notably between the segments (intersegmental membranes),
in the pleurotergal (thorax) or pleural regions (abdomen), and on the ventral sides of
the thoracic segments. The flexible parts have a higher proportion of chitin.
The hardening process transforming arthropodin into sclerotin takes place in
several steps. Prosclerotin is an intermediate product. A cuticle where the tanning
process terminates at an intermediate stage is referred to as mesocuticle. It is hardened but not fully pigmented, and can be stained with acid fuchsine (Chapman 1998).
This type occurs in transition areas between sclerites and membranes (semimembranous areas).

1.1.2 Canals and pores
The endo-, meso- and exocuticle are perforated by pore canals. Very thin cellular
processes of the hypodermis are involved in their formation during the secretion of
the cuticle. They are usually withdrawn after the process is complete. The shape of the
canals is often helical, following the arrangement of the chitin micelles in different
layers. Single epidermal cells can form numerous pore canals, up to 200 in Periplaneta americana (Seifert 1995), which is equivalent to more than a million per mm2. The
lumen can vary between ca. 15 nm (Periplaneta americana) and 100 nm. The hypodermal cytoplasmic processes are maintained in the proximal parts of the channels


4

1 Morphology

in some groups, but usually very fine chitin filaments are formed in the lumen. In the
exocuticle of sclerites they are often filled with chitin-free sclerotin, which increases
the mechanical stability in the vertical direction (Seifert 1995). At the base of the epicuticle each pore canal divides into several branches, which are filled with wax and
perforate the dense lamina and the cuticulin layer (Seifert 1995).

1.1.3 Surface structures, microtrichia and setae
The cuticle is usually characterized by fine surface patterns reflecting the arrangement of subtending hypodermal cells. A multitude of surface structures and modifications occur in different lineages. Minute spines or tubercles can be formed by the
epicuticle alone, but the exocuticle is usually involved in the formation of surface
structures. Simple surface modifications are the solid microtrichia (=trichomes),
which are not articulated and not in contact with the hypodermis after their formation. They are never associated with sensory cells and are referred to as acanthae
when they are formed by a single cell. Specialized microtrichia, usually with a soft,
flexible cuticle and a spatulate apical part, form attachment devices of the hairy type
(e.g., hairy soles of tarsomeres; see 1.3.5 Attachment structures).

set
mr

brc

toc

bri

trc

hypc

Fig. 1.1.3.1: Seta and associated cells, schematized. Abbr.: brc –
basal ring cell, bri – basal ring, hypc – hypodermal cell, mr –
membrane ring, set – seta, toc – tormogen cell, trc – trichogen cell.
Redrawn from Seifert (1995).

Setae or true hairs articulate in a flexible socket or diaphragm permitting movement
(Figs 1.1.1.1, 1.1.3.1). They are formed by a specific hypodermal cell, the trichogene cell,



1.1 Integument

5

and are always in contact with the hypodermis. The socket is formed by the tormogene
cell and has usually three layers, the external joint membrane, a ring of suspensory
fibers, and the thin and fibrous socket septum. The joint membrane is enclosed by the
cuticular basal ring. Setae are primarily hair-like but can be modified in many different ways, especially in the context of sensorial functions (see 1.1.4 Cuticular sensilla).
Cuticular scales occur in Collembola, Diplura, Archaeognatha and Zygentoma, and are
possibly an apomorphic groundplan feature of Hexapoda. The scales of Lepidoptera
and Archostemata (Coleoptera) are neoformations derived from hair-like setae. Clubshaped setae occur in 1st instar nymphs of Orthoptera and spear-shaped defensive hairs
are present in larvae of Dermestidae (Coleoptera). Long and flexible swimming hairs
occur in different lineages of aquatic beetles (e.g., Dytiscidae, Hydrophilidae) and also
in aquatic groups of Heteroptera (e.g., Corixidae, Notonectidae).

1.1.4 Cuticular sensilla
Sensilla are the basal functional and structural units of cuticular mechanoreceptors
and chemoreceptors. They include the cuticular component (e.g., the cuticle of a
seta), the sensory neuron (or neurons), the associated sheath cells with the cavities
they enclose and the structures they produce (Figs 1.1.1.1, 1.1.4.1) (Chapman 1998).
cutsh
dt

sen

set
bri

dt

thc

hypc

A

toc

trc
ax

B

thc

sec

Fig. 1.1.4.1: Cuticular sensilla, schematized. A, mechanoreceptive seta; B, chemoreceptive
sensillum. Abbr.: ax – axon, bri – basal ring, cutsh – cuticular sheath, dt – denrite, hypc –
hypodermal cell, sec – sensory cell, sen – chemoreceptive modified seta, set – mechanoreceptive
seta, thc – thecogen cell, toc – tormogen cell, trc – trichogen cell. Redrawn from Seifert (1995).


6

1 Morphology

The least modified type is the sensillum trichodeum (hair sensillum). It is composed of a primary sensory neuron and a hair-like seta with a wall consisting of exocuticle and epicuticle (Figs 1.1.1.1, 1.1.4.1). The typical mechanoreceptive seta tapers from
the base towards its apex. The displacement of the hair in the socket results in the
neural stimulus (Chapman 1998). Sensilla chaetica are shorter and thick-walled hairlike mechanoreceptors. Very long and thin sensilla not tapering apically are sometimes
referred to as trichobothria. They are not homologous with the true trichobothria
occurring in Arachnida.
The cells involved in the formation of a sensillum are derived from the same hypodermal cell (sense organ precursor cell, sense organ mother cell). In addition to the
tormogen and trichogen cells, a small thecogen cell is present between the latter and
the neuron. It secretes a cuticular layer around the distal dendrite, the dendrite sheath,
which usually ends at the base of the hair.
Modified types of cuticular sensilla are the apically rounded sensillum basiconi­
cum, and the sensillum campaniformium, which scarcely protrudes beyond the
basal ring enclosing it (Fig. 1.1.4.2). The sensillum placodeum is entirely flat and the
sensilla coeloconica and ampullacea are sunk below the external cuticular surface
to different degrees. The shape of the outer cuticular element of the sensillum does
not necessarily indicate its function. Larger hair-like sensilla are in most cases mechanoreceptors, but can also function as contact chemoreceptors. A function as olfactory
chemoreceptor is indicated by fine pores in the wall of the sensillum. Pores (apart from
the molting pore) are always absent from sensilla solely functioning as mechanoreceptors (e.g., Chapman 1998). Therefore they are also referred to as aporous sensilla.
The sensorial neurons associated with sensilla trichodea are usually of the bipolar
type (Figs 1.1.1.1, 1.1.4.1). Basally they are enclosed by the neurilemma (glial sheath).
The distal dendrite is ensheathed by the thecogen cell and is strongly narrowed where
it enters the base of the seta. Its distal part is characterized by one or two concentric
rings of microtubule doublets (9 × 2 + 0 pattern) and is consequently called the sensorial cilium. The receptive element is the membrane of its apical part, the tubular body,
which comprises a bundle of numerous microtubules (between 30 and up to 1,000)
connected by an electron-dense substance. The apical part of the tubular body is firmly
embedded in the cuticle of the base of the sensillum trichodeum. The sensorial cilium
(including the tubular body) is enclosed in the cuticular dendritic sheath, which is
secreted by the thecogen cell. In contrast to mechanoreceptors, the distal dendrites of
chemoreceptive cells do not insert into the cuticle at the base of the sensillum by way
of a tubular body. Chemoreceptors are often equipped with several sensorial cells of
different modality. Each individual dendrite can be enclosed in a tubular sheath but
often all dendrites are enclosed within a multilocular dendritic sheath with one in each
loculus. The dendrites usually branch after entering thin-walled olfactory sensilla,
which are characterized by pores arranged in hexagonal groups. The diameter usually
ranges between 15 and 20 nm. Each pore widens within the cuticle and forms a pocket.



1.1 Integument

A

B

D

E

7

C

F
Fig. 1.1.4.2: Different types of cuticular sensilla. A, sensillum trichodeum;
B, s. basiconicum; C, s. campaniformium; D, s. placodeum;
E, s. coeloconicum; F, s. ampullaceum. Redrawn from Seifert (1995).

Extremely fine pore tubules (5–6 nm) originate from the pockets and end in the liquor
of the sensillum adjacent to the dendritic membrane (Seifert 1995).

1.1.5 Scolopidia
Scolopidia are specialized internal (subcuticular) mechanoreceptors (Fig. 1.1.5.1) and
probably derived from sensilla trichodea. They consist of the scolopale cell which is
homologous to the thecogen cell, the scolopale cap cell which is equivalent to the
trichogen cell (enclosed by a tormogen cell), and one or several bipolar sensory
neurons. Scolopidia often function as proprioreceptors or are sensitive to vibrations
of air or substrates.
An external hair-like element is not present. The cell body of the sensorial cell
is sunk below the hypodermis. It is connected with it by the external scolopale
cap cell (=attachment cell), which is placed on top of the scolopale cell and
the dendrite, which is proximally covered by a sheath cell and apically narrows
to a cilium-like process containing a peripheral ring of nine microtubule doublets
with proximally extending roots. The doublets are often (or perhaps always) connected with the cell membrane near their origin at the basal body by a structure
called the ciliary necklace (Chapman 1998). The cuticular scolops is secreted by the
scolopale cell and rests on the apical part of the dendrite like a cap. The scolopale
cell contains the scolopale, which consists of fibrous material containing actin


8

1 Morphology

cut
capc
sccap

cil
roap
rocil
scolc

dend

shc
nucl
sec

ax

swc
Fig. 1.1.5.1: Scolopidium. Abbr.: ax – axon, capc – cap cell, cil –
cilium, cut – cuticle, dend – dendrite, nucl – nucleus, roap –
root apparatus, rocil – root of cilium, sccap – scolopale cap, scolc –
scolopale cell, sec – sensory neuron, shc – sheath cell, swc –
Swann cell. Redrawn from Seifert (1995), after Gray (1960).

arranged in ring or a series of rods (scolopale rods) (Chapman 1998). In scolopidia of
the subintegumental (=mononematic) type the distal end of the cap lies completely
below the body surface. In integumental (=amphinematic) scolopidia the scolops
is attached to the external cuticle by a thin cuticular terminal extending though a
narrow fold of the scolopale cap cell.
Chordotonal (or scolopophorous) organs (Fig. 1.1.5.2) are formed by scoloparia,
groups of scolopidia stretching between two movably connected sclerites. They are
specialized mechanoreceptive organs (either proprioreceptive or exteroceptive).
Examples are Johnston’s organ in the second antennomere (pedicel) of Insecta, or the
subgenual organs in the distal parts of the legs which perceive substrate vibrations.



1.1 Integument

9

flag

ped

ann

Joor
scoln

sca
Fig. 1.1.5.2: Chordotonal organ, Johnston’s organ of the pedicellus,
Melolontha vulgaris (Coleoptera, Scarabaeidae).
Abbr.: ann – antennal nerve, flag – 1st flagellomere,
Joor – Johnston’s organ, ped – pedicellus, sca – scapus,
scoln – scolopale nerve. Redrawn from Seifert (1995),
after Snodgrass (1935).

1.1.6 Integumental gland cells
Gland cells associated with the integument are embedded in the hypodermis
(Fig. 1.1.1.1: glc, see above) and much rarer than the unmodified epidermal cells surrounding them. They usually produce secretions permanently. The shape is more
rounded compared to other cells of the hypodermis and they are often extended
towards the body cavity. In the typical case the nucleus appears enlarged, irregularly
lobate or star-shaped. Endopolyploidy is common. Three types of cells with specialized glandular functions were distinguished by Noirot & Quennedey (1974). Class 1
gland cells have the apical membrane produced as microvilli or lamellae, which are in
direct contact with the cuticle secreted by themselves. They are often involved in the
production of pheromones. Class 2 (only present in termites) and class 3 cells have no
contact with the cuticle. Microvilli are present around vesicles structurally associated
with a duct forming a connection to the exterior is absent in the former but present in
the latter (Chapman 1998). The duct is an invagination of the external cuticular layer
(Seifert 1995). Microvilli are absent from class 2 cells (Noirot & Quennedey 1974).



10

1 Morphology

1.1.7 Ecdysis
The ecdysis, i.e. the molting, shedding and replacement of the cuticle, was traditionally considered as an autapomorphy of Arthropoda in the widest sense, i.e. also
including Onychophora and Tardigrada (“Panarthropoda”). This interpretation was
based on the Articulata-concept, with Annelida (“ringed worms”) and arthropods
as sistergroups. It is assumed today that it evolved earlier, as a derived groundplan
feature of a clade Ecdysozoa, which includes Arthropoda and the Cycloneuralia
(Nematoda, Nematomorpha, Priapulida, Kinorhyncha, Loricifera).
Due to mechanical properties of the cuticle the extensibility of the integument is
very limited. Therefore, before reaching their maximum size and maturity, hexapods
and other arthropods molt several times. During these intervals the integument, i.e.
the hypodermis (=epidermis) + cuticle, undergoes a period of expansion, and this
allows an increase in size of the body. However, ecdysis affects not only the body
surface, but also endoskeletal elements (tentorium, furcae, pleural ridges etc.) and
other chitinized internal invaginations such as the tracheae and also the ectodermal fore-and hindgut (see 1.9 Digestive tract). The cuticle of these structures is also
replaced during molts.
The succession of ecdyses divides the life cycle of hexapods and other arthropods
into a series of stages or instars. The number of stages in the postembryonic development differs strongly between groups and depends on different factors, such as for
instance, availability of food, temperature, or humidity. It is mainly taxon-specific
and usually relatively constant but may even vary between individuals of the same
species in some groups. Molting stops after maturity is reached in most groups of
hexapods. However, this is not the case in the basal apterygote lineages, which are
characterized by a large number of molts (e.g., up to 50 in Collembola). The number
of ecdyses is still relatively high in basal pterygote orders (e.g., Ephemeroptera,
Odonata, Plecoptera) but most insects molt only 4–6 times before reaching the adult
stage. Ephemeroptera are the only insects molting as an immature winged instar, the
subimago.
Molting comes at an evolutionary cost corresponding to the various benefits of a
solid integument (see 1.1.1 Cuticle and epidermis). Ecdyses are always critical intervals in the life cycle: hexapods and other arthropods lack their mechanical protection
during this process and their mobility is strongly restricted. The condition of hexapods just after ecdysis is called teneral.
Molting starts with the apolysis, the separation of the old cuticle from the epidermis. This is induced by an increased level of ecdysteroids functioning as molting
hormones. The size of the epidermal cells increases and a series of mitoses take place
subsequently. Shortly before and immediately after apolysis vesicles within each epidermal cell release their electron-dense contents at the cell apex. These are principally enzymes involved in the degradation of the old cuticle or material for building
the new one. The vesicles are still recognizable below the old endocuticle before they


1.1 Integument

11

release their contents. A thin hyaline and homogenous lamina formed from the inner
layers of the endocuticle is called ecdysial membrane. Due to a specific sclerotization
process it is not affected by enzymes in the following stages of ecdysis. That part of
the cuticle separating from the epidermis and the ecdysial membrane is referred to as
the exuvia and the gradually expanding space below it as ecdysial space. The latter
is filled with exuvial fluid secreted by the epidermal cells. The enzymes begin with the
degradation of the old endocuticle after a short period of inactivity. The formation of
the new cuticle is not affected due to the barrier provided by the ecdysial membrane.
The degradation products of the old cuticle are absorbed by the epidermis and used to
build the new exoskeleton in the following process (Dettner & Peters 2003).
The secretion of new cuticle starts during the degradation of the old one. This
process is initiated by the formation of the epicuticular cuticulin layer on top of the
projections or ridges of the apical membrane of the epidermal cells. After consolidation and hardening of this lamina the secretion of the new inner layers of the epicuticle and of the procuticle starts. The precise modalities of the hardening of the
external procuticle, i.e. the formation of the exocuticle, are still disputed. Phenoloxidases (tyrosinases, laccases) apparently play a role in this process, but also in other
functional contexts such as the repair of damaged cuticle or melanization (Dettner &
Peters 2003). Movements of epidermal microvilli and plaques at the apices of these
minute structures are probably responsible for the regular arrangement of chitin filaments.
The final stage of molting is the ecdysis in the narrow sense, the shedding of the
old epi- and exocuticle. In most groups it splits open at the epicranial sutures (frontaland coronal sutures) and the dorsomedian ecdysial line of the postcephalic tergites.
Muscle contractions usually increase the haemolymph pressure in the anterior body,
which results in the rupture of the old exoskeleton at the dorsal preformed zones of
weakness. The teneral exoskeleton is soft, unpigmented and wrinkled. During this
stage, the expansion of the body takes place involving locally increased haemolymph
pressure (e.g., in the limbs) and often also air uptake, especially in larger hexapods.
In the typical case the tanning process in the exocuticle results in the re-formation
of a hardened and pigmented exoskeleton within a few hours after eclosion, but this
process can also take days or even weeks in the members of some groups. After the
normal mechanical properties of the cuticle are restored, the animal has regained its
full mobility and mechanical protection.
[Snodgrass (1935); Gray (1960); Noirot & Quennedey (1974); Seifert (1995); Chapman (1998); Dettner
& Peters (2003)]



12

1 Morphology

1.2 Head
1.2.1 Segmentation, sutures and cephalic regions
The head (Figs 1.2.1.1, 1.2.2.1) is a compact and complex tagma with a concentration of
a broad array of structures und functions. It is equipped with sense organs such as the
highly complex compound eyes (Fig. 1.6.2.1) and the antennae, an elaborate set of
mouthparts (Fig. 1.2.4.1), and a complex muscle apparatus (see Table I for all cephalic
muscles). It also contains central elements of the nervous system, the anterior part of
the digestive tract including the cibarium (preoral cavity) and the salivarium, and
also neurohaemal organs (Fig. 1.11.2). It is likely that the hexapod head is homologous to a six-segmented cephalon which may represent a groundplan apomorphy
of Euarthropoda (Chelicerata, Myriapoda, Pancrustacea [=Tetraconata, crustaceans
and Hexapoda]). A six-segmented head was likely present in the extinct †Trilobita
and other fossil lineages, and is preserved in the extant Myriapoda and Pancrustacea (fused with thoracic segments in most crustacean subgroups: cephalothorax). In
contrast to †Trilobita, the primary head segmentation in hexapods cannot be traced
directly by external segmental borders, but it can be deduced from the tripartite cer­
ebrum (brain) and tripartite suboesophageal ganglion (Figs 1.5.1.1–1.5.1.3) and the
appendages, i.e. the antennae, mandibles, maxillae and labium (the segmental
affiliation of the labrum is disputed). Territories of the head were identified by different authors, but it was pointed out by Denis & Bitsch (1973) that they do not exactly
correspond with the primary segments. Unlike in crustaceans, the 3rd head segment
(=intercalary segment) of hexapods, which is associated with the tritocerebrum,
lacks appendages. The posterior three segments, associated with the three parts of
the suboesophageal ganglion and the mouthparts, are sometimes referred to as gna­
thencephalon. The “occipital ridge” was postulated as a border separating the maxillary and labial segments. However, this line often depicted in schematic drawings is
either absent or only present as a relatively short dorsolateral furrow, which does not
delimit a primary “occipital segment”.
Different head regions separated by “sutures” can be distinguished in most
hexapods (Fig. 1.2.1.1). However, it has to be noted that what is generally addressed
as “sutures” comprises two very distinctly different structural modifications of the
external cuticle (e.g., Wipfler et al. 2011):
a) Molting lines (frontal and coronal sutures [=epicranial lines or sutures]). In the
following only these will be addressed as sutures.
b) Internal strengthening ridges (e.g., clypeofrontal “suture”). In the following they
will be addressed as ridges or strengthening ridges. They are not lines of fusion or
molting lines.



1.2 Head

ce

13

occr

oc

cmem

flag
poccr
ped
moc

ge
fr

sca

sge
cly

cescl
md
lbr

pgl
plb

lac
ga

gl
pmx

Fig. 1.2.1.1: Generalized hexapod head, lateral view. Abbr.: ce – compound eye, cescl –
cervical sclerite, cly – clypeus, cmem – cervical membrane, flag – flagellum, fr – frons,
ga – galea, ge – gena, gl – glossa, lac – lacinia, lbr – labrum, md – mandible, moc –
median ocellus, oc – ocellus, occr – occipital ridge, ped – pedicellus, pgl – paraglossa, plb –
palpus labialis, pmx – palpus maxillaris, poccr – postoccipital ridge, sca –
scapus, sge – subgena. Redrawn from v. Kéler (1963).

1.2.2 Head capsule
The head capsule is almost always a rigid, well sclerotized structural unit, equipped
with a defined set of appendages (Fig. 1.2.1.1). It is reinforced by strengthening ridges
and the endoskeletal tentorium (Fig. 1.2.2.1). The shape is varying from more or
less globular (e.g., Orthoptera) (Fig. 6.12.1) to strongly flattened and elongated (e.g.,
Raphidioptera) (Fig. 6.28.1A). It is connected with the prothorax by a more or less
wide cervical membrane, often reinforced by articulatory cervical sclerites. The
posterior head region can be fully exposed (e.g., Zoraptera, Hymenoptera, Diptera)
or more or less strongly retracted into the prothorax (e.g., Blattodea, Coleoptera)
(Fig. 6.29.1). The foramen occipitale can be very narrow (e.g., Diptera), moderately
wide (most Coleoptera), or very wide (e.g., Orthoptera). It is usually divided by the
tentorial bridge into a larger upper part (alaforamen) and a smaller lower part (neu­
roforamen), and strengthened by the more or less wide postoccipital ridge, which
also serves as a muscle attachment area (Fig. 1.2.1.1).



14

1 Morphology

In hexapods with an orthognathous head (e.g., Orthoptera, Zoraptera, Hymeno­
ptera) (Figs 6.12.1, 6.13.1, 6.25.1) the mouthparts are ventrally directed, whereas they
are anteriorly directed in those with a prognathous head (e.g., Grylloblattodea,
Raphidioptera) (Fig. 6.14.2). A prognathous head is usually but not always found in
predacious hexapods. Subprognathism is an intermediate condition, with a slightly
to moderately inclined head. In the hypognathous head the mouthparts are posteriorly directed (e.g., Auchenorrhyncha, Sternorrhyncha) (Figs 6.24.1, 6.24.4). Hyperprognathism with dorsally directed mouthparts is an unusual condition occurring in
larvae of Hydrophilidae (Coleoptera).
The different head regions usually bear a characteristic vestiture of setae (articulated hairs) (Figs 6.13.2, 6.25.1). The distribution pattern of setae and pores (chaetotaxy) is taxonomically informative in some groups, especially in larvae (e.g., Lepidoptera). The coloration and surface structure of the cuticle can differ strongly in
different groups.
The compound eyes (see 1.6 Photoreceptor organs) are usually well-developed
and more or less strongly convex and round, oval or kidney-shaped. In some groups
they are small and flat (Fig. 6.14.2) or even largely (e.g., Siphonaptera) or completely
reduced (e.g., Protura, Diplura [=Nonoculata]). They are almost always placed laterally. A partial or complete subdivision occurs in few groups (e.g., males of Ephemeroptera, Gyrinidae). In Gyrinidae (Coleoptera) the upper and the lower parts are
usually widely separated and differ in their fine structure. The transparent cuticle
covering the ommatidia is thin, thus representing a zone of mechanical weakness of
the head capsule. This is usually compensated for by the presence of a broad internal circumocular ridge. Ocelli (median eyes) (Fig. 1.6.2.3) are also present in most
groups (e.g., Fig. 6.7.1). Three of them are usually arranged in a triangle on the frontal
region. They are reduced in different groups (e.g., Dermaptera, Embioptera), often in
correlation with flightlessness (e.g., Zoraptera [apterous morphs], Grylloblattodea).
The frontal and coronal (=epicranial) sutures are usually present (Figs 1.2.1.1,
1.2.2.1A). The former enclose a triangular frons in most groups (sometimes U-shaped).
Both sutures split open during the molting process. The area laterad the coronal suture
is referred to as vertex and the lateral head region as gena. However, both areas are
not delimited by a suture or ridge. A lateral area above the lateral mandibular base,
the subgena, is separated from the genal region by the subgenal ridge, which can be
divided into an anterior pleurostomal ridge above the mandibular base and a posterior hypostomal ridge. The posterior part of the subgena, traditionally assigned
as postgena, is usually not defined as a separate element. The antennal foramen is
enclosed by a circumantennal ridge, which often bears a small articulatory process,
the antennifer.
The clypeus is often trapezoid (Figs 1.2.1.1, 1.2.2.1A). Its broader posterior margin
is primarily separated from the frontal region by a transverse frontoclypeal strength­
ening ridge (=frontoclypeal “suture”, epistomal “suture”). The clypeus is sometimes
divided into an anterior, transparent anteclypeus without muscle attachment, and


1.2 Head
cors
dta

ddil

ph

poccr

poccr

ddil

tb

oes

pta

vdil
tb

ptg

lbrlev

pcly
ata

acly

A

sald

md

fr

lbr

atg

15

hyretr
sal

cdil

B

lbr
epiph

cib

plb
hyp

gl

Fig. 1.2.2.1: Generalized hexapod head, internal structures. A, anterolateral view,
frontal side of head capsule opened; B, sagittal section, brain and suboesophageal
complex removed. Abbr.: ata – anterior tentorial arm, acly – anteclypeus, atg –
anterior tentorial groove, cib – cibarium, cdil – cibarial dilator (M. clypeobuccalis),
cors – coronal suture, ddil – dorsal pharyngeal dilator (M. fronto-/verticopharyngalis),
dta – dorsal tentorial arm, epiph – epipharynx, fr – frons, gl – glossa, hyp –
hypopharynx, hyretr – hypopharyngeal retractor, lbr – labrum, lbrlev –
external labral levator (M. frontohypopharyngalis), md – mandible,
oes – oesophagus, pcly – postclypeus, ph – pharynx, plb – palpus labialis,
poccr – postoccipital ridge, pta – posterior tentorial arms, ptg –
posterior tentorial groove, sal – salivarium, sald – salivary duct, tb –
tentorial bridge, vdil – ventral pharyngeal dilator (M. tentoriopharyngalis).
Courtesy of H. Pohl, redrawn from Weber & Weidner (1974), with modifications.

a posterior sclerotized postclypeus (Fig. 1.2.2.1A), which serves as attachment area
for epipharyngeal muscles (M. clypeopalatalis3). The dorsolateral clypeal margin is
marked by the anterior tentorial pits or grooves, which represent the areas of invagination of the anterior tentorial arms.
The foramen occipitale may be narrowed by a sclerotized, unpaired gula, especially in prognathous forms (e.g., Coleoptera, Megaloptera, Raphidioptera). The gula
is likely formed by a sclerotized ventromedian region of the cervical membrane and
is usually distinctly separated from the laterally adjacent parts of the head capsule
by internal gular ridges. The landmark between the gular region and the posterior
labial (submental or postmental) margin is marked by the posterior tentorial pits or
grooves, the posterior invagination sites of the tentorium, which are generally present
in hexapods, with very few exceptions (e.g., Strepsiptera). An alternative partial closure

3 In the following names of Wipfler et al. (2011) are used for cephalic muscles. A muscle table (Table I)
with the nomenclature for head muscles is presented at the end of the chapter 1.2 Head.



16

1 Morphology

of the foramen occipitale can be formed by a hypostomal (or postgenal) bridge, i.e.
a mesally projecting duplicature of the posterolateral head capsule (e.g., Hymenoptera
partim).
Muscles not associated with movable appendages or the digestive tract are largely
or completely absent in hexapods. Exceptions are muscles linking the tentorium with
the external wall of the head capsule and muscles associated with the antennal
hearts (pulsatile organs).

1.2.3 Cephalic endoskeleton
The head endoskeleton (Fig. 1.2.2.1) comprises invaginated sclerotized elements and
also mesodermal ligamentous structures in the most basal hexapod lineages (e.g.,
Archaeognatha, Tricholepidion). Ridges play an important role as muscle attachment
areas and mechanical strengthening elements. The postoccipital ridge around the
foramen occipitale is often an attachment area for bundles of the mandibular flexor
and extensor and also for extrinsic head muscles. The circumocular ridge strengthens the zone of weakness resulting from the thin cuticle of the compound eyes and
is sometimes an area of origin of bundles of the mandibular extensor. The transverse
strengthening ridge separates the clypeus from the frons. Main elements of the tentorium of Insecta are the strongly developed posterior arms, which usually arise
immediately anterad the foramen occipitale. The invagination sites are the posterior
tentorial pits or grooves. The posterior arms are usually connected with each other
by the tentorial bridge (=corpotentorium) and also fused with the anterior tento­
rial arms in Pterygota (connected by muscles in apterygote hexapod lineages). The
tentorial bridge of Pterygota (and Maindroniidae [Zygentoma]) is a product of fusion of
transverse connecting bars of the anterior and posterior arms, respectively (see 6 The
hexapod orders [Dicondylia]). In some groups an accessory bridge is formed by medially fused internally directed processes, the laminatentoria. This feature is sometimes
erroneously referred to as “perforated corpotentorium”. It is likely an autapomorphy
of Dictyoptera but does also occur in other groups (e.g., some groups of beetles). The
anterior tentorial pits or grooves are the invagination sites of the anterior arms. They
arise at the proximolateral edge of the clypeus, close to the secondary mandibular
joint, almost always on the transverse frontoclypeal strengthening ridge (epistomal
ridge) or on the subgenal ridge. The dorsal tentorial arms originate from the anterior
arms (most groups of Polyneoptera) or at the junction area of the posterior and anterior
arms. They are not invaginations of the head capsule but usually connected to it by
fibrillar structures (fused with the wall of the head capsule in some cases).
In apterygote hexapods the tentorium is held in its position by several muscles
that originate on the head capsule. Only one muscle (M. tentoriofrontalis anterior) is
preserved in lower neopteran insects. Additionally, the posterior arms almost always
serve as attachment areas of extrinsic maxillary, labial and hypopharyngeal muscles,


1.2 Head

17

and the corpotentorium as an area of origin of ventral pharyngeal dilators. The dorsal
and anterior arms are usually attachment sites of extrinsic antennal muscles, and the
latter also of tentorial muscles of the mandibles.
Reductions and simplifications of the tentorium occur, especially but not only in
groups with reduced or strongly modified mouthparts (e.g., Strepsiptera, Diptera).

1.2.4 Labrum and epipharynx
The labrum (Figs 1.2.1.1, 1.2.2.1, 1.2.4.1), is an unpaired anterior appendage of the head
and forms the anterior closure of the preoral cavity between the paired mouthparts.
It usually covers a considerable part of the mandibles. The preoral cavity or cibar­
ium can be defined as the space between the inner labral wall (ventral4 epipharynx)

lbr
md

inc

hyp
con

pmx

molar

lac

pmt

cd

ga

susp
pgl

gl

plb

mt

sti

smt

Fig. 1.2.4.1: Mouthparts of a generalized insect (Periplaneta americana, Blattodea),
labrum (top), mandibles, hypopharynx (center), maxillae, labium (bottom). Abbr.:
cd – cardo, con – condyle (primary mandibular joint), ga – galea, gl – glossa, hyp –
hypopharynx, inc – mandibular incisivi, lbr – labrum, lac – lacinia, md – mandible,
molar – molar region, mt – mentum, pgl – paraglossa, plb – palpus labialis, pmt –
prementum, pmx – palpus maxillaris, smt – submentum, sti – stipes, susp –
hypopharyngeal suspensorium. Courtesy of H. Pohl.

4 The terms referring to directions (e.g., anterior, ventral) apply to an orthognathous head here and in
the following. The corresponding terms for a prognathous head are: anterior (orthognathous) = dorsal
(prognathous); posterior = ventral; dorsal = posterodorsal; ventral = anterior.



18

1 Morphology

(Figs 1.2.2.1A, 6.18.2B) and the inner surface of the clypeal region (dorsal epipharynx)
on one hand, and the hypopharynx on the other (see below). A wider preoral space
limited by the labrum and labium would also include the salivarium (see below).
The proximal border of the epipharynx (and cibarium) is the anatomical mouth, the
anterior opening of the pharynx.
The homology of the labrum is disputed. Its insertion below (or anterad in
pro­gnathous hexapods) the antennae suggests that it may belong to the first head
segment, possibly homologous to the antennae of Onychophora and the “great
appendages” of Cambrian fossils (e.g., †Anomalocarida). However, it was pointed out
by Denis & Bitsch (1973) that a true preantennal segment (and the “acron”) is not traceable in Hexapoda. Moreover, the innervation from the tritocerebrum and embryological evidence suggest that it may rather belong to the intercalary (= 3rd) segment.
At its base the labrum is usually moveably connected with the ventral clypeal
margin by an internal membranous fold. However, it is sometimes immobilized
and more or less completely fused with the clypeus (e.g., predacious beetle larvae:
Adephaga, Hydrophiloidea, Cleroidea etc.). The product of fusion is called clypeola­
brum or frontoclypeolabrum if the frontoclypeal strengthening ridge is absent. The
anterior labral wall is almost always sclerotized and equipped with setae. The ventral
(anterior in prognathous insects) and lateral margins are usually rounded. A more or
less deep ventromedian emargination is present in different groups (e.g., Orthoptera,
Blattodea partim). Sensilla and/or microtrichia are often present along the ventral
edge. Basolateral sclerotized rods, the tormae, reinforce the epipharynx and their
distal part serves as attachment area of the lateral labral retractors (see below).
The inner labral wall, i.e. the ventral epipharynx (Figs 1.2.2.1B, 6.18.2B), is usually
largely membranous or semimembranous. It often bears a vestiture of specifically
arranged microtrichia (non-articulated hairs), which are usually posteriorly directed,
thus facilitating the food transport towards the anatomical mouth. In some groups
of insects the lateral margin of the dorsal (clypeal) epipharyngeal part is fused with
the lateral margin of the dorsal hypopharynx, thus forming a more or less elongated
prepharyngeal tube, which is continuous with the pharynx.
M. labroepipharyngalis, which is present in most lineages, connects the anterior
and posterior labral wall. Its contraction widens the anterior region of the preoral
cavity. Other intrinsic labral muscles can occur in apterygote hexapods, ephemeropterans and in members of Polyneoptera. The labrum is retracted by two pairs of
extrinsic muscles in the groundplan of hexapods (reduced in different groups). M.
frontolabralis (Figs 6.14.3, 6.18.2B) usually originates on the median frontal region
and inserts medially on the external labral wall. M. frontoepipharyngalis originates
laterad the former and inserts on the tormae. Reduction of labral muscles occur in different groups. M. frontolabralis, for instance, is always absent in Coleoptera.
In some groups the labrum is strongly modified. In some basal lineages of Diptera
(e.g., Culicidae) and in Siphonaptera it forms a part of the piercing-sucking apparatus (Figs 6.35.1, 6.35.2) with a food channel on its ventral (epipharyngeal) side. The


1.2 Head

19

labrum partly covers the labial rostrum in Hemiptera (Figs 6.21.2, 6.24.1, 6.24.4). It is
almost always completely reduced in Strepsiptera but still present in the groundplan
of the order.

1.2.5 Antennae
The antennae are the paired appendages of the 2nd segment and homologous to the
1st antennae (=antennulae) of crustaceans (Figs 6.1.1, 6.1.2, 6.4.1). The antennal nerve
originates from the second part of the brain, the deutocerebrum. In Collembola and
Diplura (and in Myriapoda and crustaceans) muscles are present in all antennomeres
(often referred to as antennal segments) except for the apical one. This type of antenna
(=“Gliederantenne” in German) belongs to the groundplan of Hexapoda (Figs 6.1.1,
6.3.3) An autapomorphy of Insecta is the antenna of the flagellar type (“Geißelantenne”) (Figs 6.4.1, 6.4.2), with muscles only in the basal antennomere, the scapus
(Mm. scapopedicellares lateralis and medialis). A chordotonal organ is present in
the pedicellus (2nd antennomere), which bears a more or less elongated flagellum
without muscles, which is usually composed of many antennomeres (flagellomeres).
The antennae are inserted in the antennal foramen (see above), between or above
the compound eyes in most groups (Fig. 1.2.1.1). They are almost always movable in all
directions and more or less densely covered with setae and different sensilla, mostly
chemo- and mechanoreceptors (Figs 4.4.1B, 4.9.1B). The scapus usually articulates
with the antennifer and is moved by three or four extrinsic antennal muscles in most
groups (Mm. tentorioscapales anterior, posterior, lateralis and medialis). The two
intrinsic muscles (see above) are attached to the base of the pedicellus and move the
remaining part of the antenna. The Johnston’s organ in the pedicellus of Insecta is a
complex chordotonal organ composed of numerous scolopidial sensilla. It functions
as a mechanoreceptor and registers movements of the flagellum. It also perceives
acoustic signals in some groups (e.g., Culicidae, Chironomidae).
The antennae can be modified in many different ways (e.g., Fig. 6.35.4). They are
completely absent in Protura (Figs 6.2.1, 6.2.2), largely reduced in larvae of some groups
of Holometabola (e.g., Strepsiptera) (Fig. 6.30.1), and shortened and more or less
bristle-like in Odonata (Fig. 6.7.1A), Ephemeroptera (Fig. 6.6.2) and Auchenorrhyncha
(Fig. 6.21.2). The ancestral type (groundplan of Insecta) is the filiform antenna with
a multisegmented, slender flagellum. In moniliform antennae the flagellomeres are
more or less globular (e.g., Isoptera, Embioptera partim), whereas they are sawtoothshaped in the serrate type (e.g., Elateridae [Coleoptera]). The flagellomeres of pectinate antennae bear elongate extensions either on one or on both sides (e.g., different
groups of Lepidoptera, Tipulidae [Diptera]). In capitate or clubbed antennae one or
several apical segments are symmetrically or asymmetrically widened (e.g., different
groups of Lepidoptera and Coleoptera). Geniculate antennae are hinged or bent (e.g.,
different groups of Hymenoptera [e.g., ants], Curculionidae [Coleoptera]). In “higher”


20

1 Morphology

dipterans (Cyclorrhapha) the first flagellomere (postpedicellus) is strongly enlarged
whereas the remaining flagellum forms a bristle-like arista (aristate type) (Fig. 6.35.4).

1.2.6 Mandibles
The mandibles are the paired appendages of the 4th head segment (Figs 1.2.1.1, 1.2.4.1)
and receive their innervation from the anterior part of the suboesophageal ganglion.
They play a dominant role in the mechanical processing of food. Unlike the maxillae
(Fig. 1.2.4.1) they are primarily a compact, undivided and strongly sclerotized element
without appendages. In Dicondylia (Zygentoma + Pterygota) they are connected with
the head capsule by a posterior primary mandibular joint (condyle on the mandible)
and an anterior secondary joint (condyle at the clypeal base). The movements of the
dicondylous mandible are restricted to a single level perpendicular to the axis between
these two articulations. The mandible is almost always distinctly curved inwards,
especially at its outer margin, and it is usually broadest at its base. One or several
apical or subapical teeth (incisivi) (Fig. 1.2.4.1) are used to perforate and crush food
substrate. Often, especially in hexapods feeding on plant matter or fungi, a mesally
directed prominence is present at the mandibular base, the mola; usually its surface
is modified for efficient grinding, with parallel ridges or densely arranged tubercles.
In predacious forms (e.g., Odonata, Mantophasmatodea) the mesal surfaces of the
mandibles form distinct cutting edges. An articulated mesal appendage, the lacinia
mobilis or prostheca, occurs in several groups. It can be sclerotized or largely membranous and is often equipped with microtrichia.
In neopteran insects with well-developed mandibles a pair of antagonistic
muscles with a cranial origin move the mandibles, the very large M. craniomandi­
bularis internus (flexor=adductor) and the distinctly smaller M. craniomandibularis
externus (extensor=abductor) (Figs 6.14.3, 6.18.2B). Both insert on strongly developed
tendons attached to the mandibular base. In the groundplan of Hexapoda strongly
developed muscles arising from the hypopharynx (M. hypopharyngomandibularis)
or the anterior tentorial arms (Mm. tentoriomandibulares lateralis superior and inferior, Mm. tentoriomandibulares medialis superior and inferior) are present, and also a
transverse muscle medially connected by a ligament (e.g., Zygentoma). They insert on
the internal surface of the mandible. The hypopharyngeal and tentorial muscles persisting in many pterygote groups are almost always very thin. One of them is usually
accompanied by a nerve and functions as a proprioreceptor.
Only the primary joint is present in Collembola, Protura and Archaeognatha
and both joints are missing in Diplura. Mandibular sucking channels occur in larvae
of some groups of beetles (e.g., Dytiscidae, Lampyridae). The mandibles are transformed into stylets in Hemiptera and biting flies (in addition to stylets formed by the
laciniae, see below). The mandibles form a sucking apparatus together with the laciniae in neuropteran larvae.


1.2 Head

21

1.2.7 Maxillae
The maxilla (Figs 1.2.4.1, 6.14.2, 6.27.3C), which is homologous to the maxillula
(1st maxilla) of crustaceans, is usually composed of cardo, stipes, lacinia (=inner
endite lobe), galea (=outer endite lobe), and palp. The relatively short cardo articulates with the head capsule, usually in a more or less deep maxillary groove (=fossa
maxillaris). At its base it bears a lateral process for muscle attachment. The articulation point lies between the insertion of the extensor and the flexor, which originate on the lateral head capsule and the posterior tentorial arm, respectively. The
stipes is connected with the cardo by a hinge. It is usually the largest part of the
maxilla and bears the palp and endite lobes. The ventral and lateral sides are sclerotized and divided into a basistipes and mediostipes in some groups (e.g., Coleoptera). The lacinia is usually more or less firmly connected or fused with the stipes
mesally. It is sclerotized and apically curved and pointed in most groups, and usually
set with strong setae or spines along its mesal edge. In groups with sucking-piercing
mouthparts (e.g., Anoplura, Hemiptera, biting flies) the lacinia is transformed into a
piercing stylet (Figs 6.35.1, 6.35.2). The galea is usually less strongly sclerotized and
equipped with chemoreceptors; it is composed of one or two segments. The maxillary
palp is usually five-segmented and also equipped with chemoreceptive sensilla; its
apex often bears a dense field of sensorial structures.
In most groups two extrinsic maxillary muscles originate from the head capsule
(M. craniocardinalis and M. craniolacinialis) and three from the posterior tentorial
arms (M. tentoriocardinalis, Mm. tentoriostipitales anterior and posterior). Additional
muscles insert on the endite lobes, on the base of the palp, and on the base of the
palpomeres.

1.2.8 Labium
The labium or lower lip (Figs 1.2.4.1, 6.14.2, 6.27.3C) forms the posterior (orthognathous head) or ventral (prognathous head) closure of the preoral space and of the
salivarium. It is homologous to the maxilla (2nd maxilla) of crustaceans, but its proximal parts form a structural unit without a recognizable median line or suture. The
proximal postmentum primarily forms the posterior closure of the head capsule
and is adjacent with the foramen occipitale unless a gula or hypostomal bridge is
present; it is often divided into a mentum and submentum (Fig. 1.2.4.1). The anterior element, the prementum, bears the appendages, i.e. the palps and the para­
glossae and glossae (outer and inner endite lobes; serially homologous to the galea
and lacinia); it may or may not be incised medially. The labial palp is usually threesegmented but otherwise very similar to the maxillary palp; it is either inserted on a
more or less distinct palpiger or directly on the distolateral edge of the prementum.
The musculature is similar to the muscle equipment of the maxilla. Three pairs of


22

1 Morphology

premental retractors are usually present in Pterygota, one of them originating on the
postlabium (M. submentopraementalis) and two on the posterior tentorial arms (Mm.
tentoriopraementales inferior and superior). In addition intrinsic premental muscles
are associated with the glossae and paraglossae, the palps, and the salivarium. Intrinsic palp muscles are also usually present.
The labium is transformed into a rostrum in Hemiptera (Figs 6.24.1, 6.24.4A). It
forms a sheath for the stylet-like mandibles and laciniae. The outer and inner endite
lobes of the labium are distinctly reduced or missing in holometabolous insects with
the notable exception of Hymenoptera. In bees, the labium forms a complex with
the maxillae. Both elements of this maxillolabial complex are connected by a small
rod-like sclerite, the lorum. The frontally fused, strongly elongated and pubescent
glossae serve as uptake apparatus of liquid food.

1.2.9 Hypopharynx
The hypopharynx is an unpaired tongue-like structure between the paired mouthparts (Figs 1.2.4.1, 6.18.2B). It is likely a derivative of several segments and not serially
homologous to the primarily paired appendages (Denis & Bitsch 1973).
The hypopharynx is usually largely semimembranous, but almost always reinforced by sclerites (Fig. 1.2.4.1: susp). It forms the posterior floor of the cibarium and
a ramp for transporting food towards the anatomical mouth (Fig. 6.18.2B). Its anterior
surface is often at least partly sclerotized and bears a vestiture of posteriorly directed
spinulae which facilitate this process (see epipharynx). Between the hypopharynx
and the anterior side of the distal part of the labium lies the salivarium, a pocket
where the salivary ducts open (see 1.2.10 Salivarium) (Fig. 6.18.2B). Superlinguae
are lateral lobes of the hypopharynx (Fig. 6.6.5), flanking the median lingua. They
are likely a groundplan feature of Hexapoda (present in Collembola, Diplura and
Archaeognatha) but they are missing in Zygentoma and distinctly reduced or absent
in Pterygota (with the exception of Ephemeroptera: Fig. 6.6.5). Reduced superlinguae
are usually represented by lateral sclerotizations, the basal sclerites or basal plates,
which sometimes extend to the salivarium. They serve as attachment areas of the tentorial retractors of the hypopharynx, M. tentoriohypopharyngalis. The suspensoria
(=fulturae) are paired sclerites or groups of sclerites embedded in the proximolateral hypopharyngeal wall (Fig. 1.2.4.1: susp). They form long anterior projections, the
oral arms which end laterad the anatomical mouth and serve as attachment area of
M. oralis transversalis, M. frontooralis and M. tentoriooralis. The last two muscles
were addressed as retractors of the mouth angle by Snodgrass (1935). Their antagonist
is M. craniohypopharyngalis, a retractor originating from the head capsule (apterygote lineages) or posterior tentorium (Pterygota). The muscle is missing in different
groups, especially in correlation with a reduced salivarium (e.g., Coleoptera). Several
other muscles linked with the hypopharynx occur in different lineages.


1.2 Head

23

The hypopharynx can be distinctly modified or reduced in different groups of
hexapods. It is greatly reduced in size in Hymenoptera and fused with the distal part
of the labium in Coleoptera, which consequently lack a salivarium. It forms one of
the stylets in basal dipteran groups (e.g., Culicidae). In Hemipterans it is perforated
by the canal of the salivary pump and continuous with the food canal of the laciniae.

1.2.10 Salivarium
The salivarium is the pocket between the posterior side of the hypopharynx and the
anterior (internal) side of the distal part of the labium (Figs 1.2.2.2B, 6.18.2B). It is
part of the preoral cavity in the wider sense and receives the openings of the salivary
ducts. The salivary glands (Fig. 1.2.10.1) are usually well developed and located in
the anterior part of the thorax. In some groups they are connected with a large reservoir (Fig. 1.9.2). The salivarium is usually equipped with three pairs of short muscles
originating from the hypopharynx (M. hypopharyngosalivaris) and the prementum
(M. praementosalivaris anterior and posterior), respectively. In certain groups a small
ring muscle (M. annularis salivarii) is present where the salivary duct opens into the
salivarium. Reductions of the salivarium (including muscles and glands) occur in different groups, notably in Coleoptera (completely absent) and in Megaloptera (vestigial).
acin

res

sald

Fig. 1.2.10.1: Salivary gland. Abbr.: acin – acini (berry-like substructure) of
salivary gland, res – reservoir, sald – salivary duct. Redrawn from Seifert (1995).

[Snodgrass (1935); Weber (1933, 1938); v. Kéler (1963); Matsuda (1965); Denis & Bitsch (1973);
Weber & Weidner (1974); Chapman (1998); Wipfler et al. (2011)]



Table I: Generalized nomenclature for cephalic muscles (based on Wipfler et al. [2011] and von Kéler [1963], modified). The terms used for the orientation of
structures refer to a prognathous head. For orthognathous heads: dorsal = anterior, ventral = posterior, anterior = ventral, posterior = dorsal.
Origin
anterior or dorsal tentorial arms
anterior or dorsal tentorial arms
anterior or dorsal tentorial arms
anterior or dorsal tentorial arms
frons
posterior or posterolateral area of scapus
mesal or posteromesal area of scapus
base of pedicellus
antennal ampulla
antennal ampulla
antennal ampulla
antennal ampulla
frons (laterally of antennal ampulla)
frons (laterally of antennal ampulla)
central region of frons or rarely on antennal base
central region of frons or rarely on antennal base
epistomal sulcus

Abbrev. Name

Antennal muscles
0an1
M. tentorioscapalis anterior
0an2
M. tentorioscapalis posterior
0an3
M. tentorioscapalis lateralis
0an4
M. tentorioscapalis medialis
0an5
M. frontopedicellaris
0an6
M. scapopedicellaris lateralis
0an7
M. scapopedicellaris medialis
0an8
M. intraflagellaris

Antennal heart muscles
0ah1
M. interampullaris
0ah2
M. ampulloaortica
0ah3
M. ampullopharyngalis
0ah4
M. ampullofrontalis
0ah5
M. frontopharyngalis
0ah6
M. frontofrontalis

Labral muscles
0lb1
M. frontolabralis
0lb2
M. frontoepipharyngalis

M. epistoepipharyngalis

M. labralis transversalis
M. labroepipharyngalis
M. labrolabralis

0lb3

0lb4
0lb5
0lb6

ventrolateral edge of labrum
centrally on labrum
centrally on labrum (between bundles of 0lb5)

1 Morphology

posterior labral margin
posterolateral edge of labrum (labro-epipharyngeal
border) or tormae
posterolateral edge of labrum (labro-epipharyngeal
border)
opposite side of ventrolateral edge of labrum
epipharynx
medially on apical labral area

antennal ampulla of opposite side
cephalic aorta
anterior pharynx
frons
dorsolaterally on pharynx
opposite side of frons

anteriorly on base of scapus
posteriorly on base of scapus
lateral margin of scapus
mesal margin of scapus
lateral edge of pedicellus
posteriorly or posterolaterally on base of pedicellus
mesal, anterior or anteromesal base of pedicellus
close to tip of flagellum

Insertion

24





M. craniomandibularis externus
anterior
M. craniomandibularis externus
posterior
M. hypopharyngomandibularis
M. tentoriomandibularis lateralis
superior
M. tentoriomandibularis lateralis
inferior
M. tentoriomandibularis medialis
superior
M. tentoriomandibularis medialis
inferior

M. stipitogalealis
M. stipitopalpalis externus

M. stipitopalpalis medialis
M. stipitopalpalis internus

0mx7
0mx8

0mx9
0mx10

Maxillary muscles
0mx1
M. craniocardinalis
0mx2
M. craniolacinialis
0mx3
M. tentoriocardinalis
0mx4
M. tentoriostipitalis anterior
0mx5
M. tentoriostipitalis posterior
0mx6
M. stipitolacinialis

0md8

0md7

0md6

0md4
0md5

0md3

0md2

Mandibular muscles
0md1
M. craniomandibularis internus

gena and postgena
posterior part of gena
tentorium
tentorium
tentorium
proximolaterally on stipes (dorsally of maxillary
palp)
laterally on stipes (dorsally of maxillary palp)
laterally on stipes (dorsally of maxillary palp) or
stipital ridge
stipital ridge
stipital ridge

anterior tentorial arm

anterior and dorsal tentorial arm

anterior tentorial arm

process of loral arm or separate sclerite close to it
anterior and dorsal tentorial arm

gena and postgena

Ventral, posterior, lateral and/or dorsal parts of the
head capsule and/or posterior and anterior
tentorial arms
gena (below compound eyes)

posteriorly on base of first maxillary palpomere
mesally on base of first maxillary palpomere

proximal part of cardo (cardinal process)
proximal egde of lacinia
distal part of cardo (close to stipitocardinal suture)
stipital ridge
proximal rim of stipes (close to stipito-cardinal sulcus)
base of lacinia (in some cases common tendon with
0mx4)
base of galea
laterally on base of first maxillary palpomere

dorsal part of mandibular base (inside of mandible)

dorsomesal area of mandible (inside of mandible)

dorsolateral area of mandible (inside of mandible)
laterally on mandibular base (between processus paratentorialis and posterior mandibular articuation
ventrally on mandibular rim (inside of mandible)

base of mandible (behind anterior mandibular articulation)
abductor (lateral) tendon of mandible

adductor (mesal) tendon of mandible

1.2 Head

25

Origin

M. tentorioglandularis
M. submentopraementalis

M. postmentomembranus
M. submentomentalis
M. praementoparaglossalis
M. praementoglossalis
M. praementopalpalis internus
M. praementopalpalis externus

M. praementomembranus
M. palpopalpalis labii primus
M. palpopalpalis labii secundus

0la7
0la8

0la9
0la10
0la11
0la12
0la13
0la14

0la15
0la16
0la17

postoccipital phragma
postoccipital phragma (laterad 0la1)
postoccipital phragma (laterad 0la2)
postoccipital phragma (laterad 0la3)
posterior tentorial arm and/or cranium
posterior tentorial arm and/or postoccipital ridge
and/or cranium
posterior tentorial arm
central or proximolateral region of submentum
or gula
postmentum
submentum
proximal rim of prementum (lateral half)
central region of prementum
distal area of prementum (medially)
medially on proximal rim or proximolateral corner of
prementum
postmentum
proximomesally on first labial palpomere
lateral wall of second labial palpomere

lateral wall of proximal part of stipes (close to
stipitocardinal suture)
M. palpopalpalis maxillae primus
laterally on base of first maxillary palpomere
M. palpopalpalis maxillae secundus mesally on base of second maxillary palpomere
M. palpopalpalis maxillae tertius
base of third maxillary palpomere
M. palpopalpalis maxillae quartus
mesally on base of fourth maxillary palpomere

M. stipitalis transversalis

Labial muscles
0la1
M. postoccipitoglossalis medialis
0la2
M. postoccipitoglossalis lateralis
0la3
M. postoccipitoparaglossalis
0la4
M. postoccipitopraementalis
0la5
M. tentoriopraementalis
0la6
M. tentorioparaglossalis

0mx12
0mx13
0mx14
0mx15

0mx11

Abbrev. Name

internal (ventral) membrane of labium
mesally on base of second labial palpomere
mesally on base of third labial palpomere

labial gland
medially on proximal premental margin or distal part
(close to galea)
internal (ventral) membrane of labium
mentum
proximolaterally on paraglossa
mesal wall of glossa
mesally on first palpomere
laterally on first palpomere

glossa
dorsolaterally on base of glossa
base of paraglossa
posterolateral corner of prementum
proximal rim of prementum
paraglossae (close to base of labial palp)

laterally on base of second maxillary palpomere
mesally on base of third maxillary palpomere
mesally on base of fourth maxillary palpomere
mesally on base of fifth maxillary palpomere

stipital ridge

Insertion

26
1 Morphology




postclypeus
postclypeus (posterad 0ci1)
frons (behind epistomal sulcus)

Cibarial muscles
0ci1
M. clypeopalatalis

Buccal muscles
0bu1
M. clypeobuccalis

M. frontobuccalis anterior

M. frontobuccalis posterior

0bu2

0bu3

posterior part of frons

lateral edge of anterior part of corpotentorium
dorsally on anterior part of corpotentorium
dorsal tentorial arm
dorsolaterally on metatentorium
ventrolaterally on anterior part of corpotentorium
anterior part of tentorium

M. craniohypopharyngalis
M. postoccipitalohypopharyngalis
M. tentoriosuspensorialis
M. postmentoloralis
M. praementosalivarialis anterior
M. praementosalivarialis posterior
M. oralis transversalis
M. loroloralis
M. lorosalivarialis
M. hypopharyngosalivarialis
M. annularis salivarii

frons
anterior tentorial arm or epistomal sulcus or frons
close to it
posterior tentorial arm or corpotentorium
postoccipital phragma
anterior margin of corpotentorium
postmentum
distolateral area of prementum (close to labial palp)
proximal or proximolateral part of prementum
oral arm of suspensorial sclerite
loral arm of suspensorial sclerite
hypopharyngeal suspensorial sclerite
hypopharyngeal suspensorial sclerite or tentorium

Tentorial muscles
0te1
M. tentoriofrontalis posterior
0te2
M. tentoriofrontalis anterior
0te3
M. tentoriofrontalis dorsalis
0te4
M. posterotentorialis
0te5
M. tentoriotentorialis longus
0te6
M. tentoriotentorialis brevis

0hy3
0hy4
0hy5
0hy6
0hy7
0hy8
0hy9
0hy10
0hy11
0hy12
0hy13

Hypopharyngeal muscles
0hy1
M. frontooralis
0hy2
M. tentoriooralis

dorsal wall of bucca (directly in front of brain)

roof of bucca (between anatomical mouth and frontal
ganglion)
dorsal wall of bucca (directly behind frontal ganglion)

roof of cibarium

frons
frons
frons
external rim of tentorium
mesally on posterior tentorial part
posterior part of tentorium

suprasalivarial sclerite
hypopharyngeal phragma
hypopharyngeal suspensorial sclerite
loral arm of suspensorial sclerite
laterally on salivarium
laterally on salivarium
oral arm of suspensorial sclerite of opposite side
oral arm of suspensorial sclerite of opposite side
suprasalivarial sclerite
salivary orifice
ring muscle close to salivary orifice

oral arm of suspensorial sclerite
oral arm of suspensorial sclerite

1.2 Head

27

dorsal tentorial arm
anterior tentorial arm or corpotentorium
posterior tentorial arm or corpotentorium

0bu4
0bu5
0bu6

posterior tentorial arm
postocciput (close to median line)

0ph2
0ph3

Stomodaeal muscles
0st1
M. annularis stomadaei
0st2
M. longitudinalis stomadaei

M. tentoriopharyngalis
M. postoccipitopharyngalis

vertex or occipitale

Pharyngeal muscles
0ph1
M. verticopharyngalis

M. tentoriobuccalis lateralis
M. tentoriobuccalis anterior
M. tentoriobuccalis posterior

Origin

Abbrev. Name

ring muscle layer covering pharynx
longitudinal muscles along pharynx

dorsally on postcerebral pharynx (directly posterad of
brain)
ventrally on postcerebral pharynx (beneath 0ph1)
posterior pharynx

lateral wall of bucca
ventral wall of bucca (directly behind anatomical mouth)
ventral wall of bucca (directly in front of brain, ventrally
of 0bu3)

Insertion

28
1 Morphology





1.3 Thorax

29

1.3 Thorax
1.3.1 Segmentation and composition of segments
The presence of a thorax composed of three clearly defined segments is arguably the
most important autapomorphy of Hexapoda. It is usually the second largest tagma of
the body and structures related to locomotion are concentrated in this region. On each
of the three segments a pair of legs is inserted ventrolaterally. Two pairs of wings with
a dorsolateral articulation area are present in most lineages of Pterygota (groundplan
autapomorphy). The three segments are more or less similar and relatively simple
in the apterygote lineages. In pterygote insects the mesothorax and metathorax
(together forming the pterothorax) are strongly modified and much more complex
than the prothorax, especially on the dorsal side (tergal region) and dorsolaterally
(wing base).
All three segments are composed of dorsal, lateral and ventral elements, the
tergum, pleuron and sternum (Figs 1.3.3.1, 6.13.3, 6.18.3). The individual regions
of sclerotized cuticle (tergites, pleurites and sternites) are connected by membranes and semimembranous areas. The membranous areas are exposed to varying
degrees in almost all groups (Fig. 6.13.3), especially on the ventral side (not in Coleo­
ptera [Figs 6.29.4, 6.29.5] and some groups of Heteroptera). The unsclerotized regions
ensure the necessary flexibility within the segments, especially at the articulation
areas of the locomotor organs. The three segments are connected by intersegmental
membranes, which are often more or less concealed. On the dorsal and ventral side
the membranous connection does not correspond with the true segmental border
(see below). Laterally the thoracic spiracles are embedded in the intersegmental
membranes (Fig. 1.3.3.1A), in most groups two pairs belonging to the meso- and metathorax, each pair located in front of the respective segment. Thoracic spiracles are
usually larger than their abdominal equivalents, which are serially homologous. The
two pterothoracic segments often form a more or less rigid functional unit, whereas
the flexibility between the pro- and mesothorax is usually relatively high. Posteriorly the metathorax is broadly connected with the abdomen in most groups. Different types of fusion of metathoracic and abdominal elements occur in various groups
(e.g., Hymenoptera).
The prothoracic tergum or pronotum is usually a simple, plate-like structure,
whereas the dorsal and dorsolateral regions of the pterothorax of pterygote insects
are modified in a complex way in correlation with the flight function (see 1.3.3 ptero­
thoracic segments) (Figs 1.3.3.1, 6.13.3). Dorsolateral tergal duplicatures (paranota,
laterotergites) probably belong to the groundplan of Insecta (preserved in the original form in Archaeognatha and Zygentoma). They are possibly precursors of the mesoand metathoracic wings of Pterygota. The lateral sclerotized element of the thoracic
segments, the pleurite, is located between the tergum and sternum. It is possibly
formed by the subcoxa, an element supposedly separating from the coxal primordium


30

1 Morphology

in the embryonic development (Matsuda 1970), but this interpretation is uncertain.
It is divided by a slanted dorsoventral pleural ridge, which is always present in the
meso- and metathorax, but often reduced or absent in the prothorax (Figs 1.3.3.1A,
6.13.3). It stabilizes the lateral body wall and divides it in the anterior episternum and
the posterior epimeron. It is also important in the functional context of the articulations of the coxae and wings. It connects the pleural wing joint with the pleurocoxal joint, the latter generally formed by a condyle at the posteroventral edge of the
ridge and a corresponding coxal concavity. An additional coxal articulation is formed
by the triangular anteroventral trochantin (Figs 1.3.3.1A, 6.13.3A), an additional
pleural element below the episternum (small and crescent-shaped, reduced in some
groups of Holometabola). The sternal region usually contains more extensive unsclerotized areas than the lateral and dorsal walls of the segments (Figs 1.3.3.1C, 6.13.3B).
However, it is almost always reinforced by several sternites. A series of five elements
is arguably present in the groundplan of Hexapoda, the short anterior presternum,
the extensive basisternum, the furcasternum (or sternellum), the spinasternum,
and the posterior poststernite (Matsuda 1970). In Pterygota, the presternum is almost
generally absent (possibly represented by isolated sclerites in Plecoptera) and the spinasternum is reduced to varying degrees in many lineages. In Holometabola the true
sternal elements are largely invaginated, mostly represented by an internal median
longitudinal ridge (discrimen), and externally largely replaced by pleural elements
(preepisternum, katepisternum).
More or less extensive ligamentous endoskeletal elements occur in the apterygote lineages. They are of subepidermal origin, usually branched, and serve as insertion points of muscles and tendons (Matsuda 1970). These structures are completely
absent in most groups of Pterygota, at least in the adult stage. The pterygote endoskeletal elements are the furcae and the smaller spinae (Fig. 1.3.3.1D). The former arise
from the furcasternum. Primarily their invagination sites are distinctly separated and
connected by a transverse ridge, the sternocosta. In correlation with a narrowed or
invaginated sternum (Holometabola) the furcal bases can be more or less closely adjacent or even arising from a common invagination site. The dorsolaterally extending
furcal arms are often fused to the pleural ridge or connected with it by short muscles
or fibrillae. They serve as muscle attachment area, especially for muscles attached
to basal elements of the legs, the coxa and the trochanter, but also for the ventral
intersegmental muscles. The spina is an unpaired invagination of the spinasternum
and serves as attachment area of coxal and intersegmental muscles. It is partly or
completely reduced in many groups, especially in the metathorax (distinctly developed in Grylloblattodea).





31

1.3 Thorax

scl

sc

acrt

pn

prsc

phr
prsc

pwp

sc

sa

pascl

anp

plr

ba

spi

scl

pnp

ep

es

pcj
tin

wilig

B

cx

phr

A
eust
eust1

fu1
fst1
sst1

fst1
spn1

eust2

cerv
eust1

fin
leg1

sst1

gul

sst

fst

pn

fin
fu2

fst2

eust2
fst2

leg2
sst2
fst3

C

spn2

eust3
fin
leg3

fst3

D

eust3
fu3

Fig. 1.3.3.1: Thoracic skeleton. A, pterothoracic segment, lateral view, schematized; B,
pterothoracic tergal region, schematized; C, sternal region, ventral view Isoperla sp.
(Plecoptera, Perlodidae); D, sternal region, internal view, Periplaneta americana (Blattodea,
Blattidae). Abbr.: acrt – acrotergite, anp – anterior notal wing process, ba – basalare, cerv –
cervical sclerite, cx – coxa, ep – epimeron, es – episternum, eust1–3 – pro-, meso-, metathoracic
eusternum, fin – furcal invagination site, fst1–3 – pro-, meso-, metathoracic furcasternum (sternellum), fu1–3 – pro-/ meso-/ metafurca, gul – gularia, leg1–3 – fore-/ mid-/ hindleg, pascl – prealar
sclerite, pcj – pleurocoxal joint, phr – phragma, plr – pleural ridge, pn – postnotum, pnp –
posterior notal wing process, prsc – prescutum, pwp – pleural wing process, sa – subalare,
sc – scutum, scl – scutellum, spi – spiracle, spn1/2 – pro-/ mesospina, sst1/2 – pro-,
mesothoracic spinasternum, tin – trochantin, wilig – wing ligament. Redrawn from Seifert (1995).



32

1 Morphology

1.3.2 Prothorax
The prothorax is anteriorly connected with the head by the cervical membrane,
usually with embedded cervical sclerites (Figs 1.3.3.1D, 6.13.3A) forming articulations and serving as muscle attachment areas. One or two pairs of lateral cervical
sclerites (laterocervicalia) are present in most groups. Unpaired ventral sclerites
occurring occasionally are called gularia. Unpaired dorsal sclerites occur in several
groups (e.g., Zoraptera, Coleoptera [Hydrophilidae]). Paired dorsal cervical sclerites
are present in Dermaptera.
The pronotum is often divided by a median line or zone of weakness (ecdysial
line in immatures) but otherwise forms a single, more or less plate-like structure, with
or without a distinct lateral edge. It mainly serves as attachment area of large dor­
soventral leg muscles and some of the extrinsic muscles of the head. The size of the
pronotum varies greatly. It forms a conspicuous pronotal shield in Blattodea, Coleo­
ptera and some other groups. In orthopterans it is saddle shaped and laterally covers
extensive parts of the pleuron. Anteriorly the pronotum covers the posterior margin of
the head in many groups. Posteriorly it often overlaps with the anterior region of the
mesotergum. Distinctly developed lateral duplicatures are present in the primarily
wingless Archaeognatha and Zygentoma (see above: paranota). Prothoracic winglets
were present in some Paleozoic insects (e.g., †Palaeodictyoptera) but absent in all
extant pterygote lineages. The prothoracic pleuron is generally less complex than the
corresponding regions of the pterothoracic segments. All modifications or structures
related to flight are lacking (e.g., basalare and subalare) (Fig. 6.13.3A). The pleural
suture is often short, indistinct or lacking. In some groups more or less extensive
propleural parts are invaginated below the pronotum, thus forming a cryptopleu­
ron (e.g., Coleoptera, especially Polyphaga). Compared to the tergal and pleural elements, the prosternum differs less profoundly from its pterothoracic counterparts,
as it is not directly connected to the flight organs. Modifications occur in different
groups, for instance in relation with hypognathous heads or raptorial forelegs. The
prosternum is often reduced in width in Holometabola but is broad in Megaloptera
and Raphidioptera.
The prothoracic musculature differs distinctly from the pterothoracic muscle
systems (Figs 1.3.3.2, 1.3.3.3, 6.13.4, see also Table II). A complex array of extrinsic
head muscles (cervical muscles, musculi cranii) is always present, whereas muscles
related to the flight apparatus are absent for obvious reasons. The leg muscles are
similar to those of the pterothorax (see below). The cervical muscles (Fig. 6.13.4;
Matsuda 1970: fig. 19, 22; Friedrich & Beutel 2008) form a complex system of dorsal
and ventral longitudinal retractors and of oblique muscles, thus guaranteeing the
movability of the head in all directions (Figs 1.3.3.2, 1.3.3.3C). They originate on the
prothoracic phragma, different regions of the pronotum, the propleuron, the pro­
sternum, and the profurca. They insert on the cervical sclerites, the tentorium and on
different regions of the posterior head capsule, especially on the postoccipital ridge.




1.3 Thorax

33

1.3.3 Pterothoracic segments
The meso- and metathorax are largely unmodified and similar to the prothorax in the
apterygote lineages (e.g., Fig. 6.1.1). Far-reaching modifications have resulted from
the acquisition of wings and functions related to flight, especially of the tergal and
pleural elements (Figs 1.3.3.1–1.3.3.3, 6.13.3, 6.13.4).
The meso- and metaterga of pterygote insects are complex and specifically subdivided structures (Fig. 1.3.3.1B). The main division is into the large anterior notum and
the transverse posterior postnotum. The notum is primarily subdivided into the short
prescutum, the extensive scutum, and a scutellum, which is usually triangular. The
part of the notum connected with the wing base is called the alinotum. In apterygote hexapods an acrotergite is separated from the prescutum by the antecosta, an
intersegmental furrow. The acrotergite and antecosta are equivalent with the postnotum of the preceding segment of Pterygota. The antecosta (or internal part of the
postnotum) forms a phragma for attachment of the dorsal intersegmental muscles.
In most groups of Pterygota it is an extensive attachment site of large dorsal indirect flight muscles and plays an important role in the flight apparatus. The transverse
prescutum is strongly narrowed in some groups. Laterally it forms the prealare (or
prealar sclerite), an attachment area of short direct flight muscles. In some groups
a process of the prescutum is bent downwards and linked with the episternum, thus
forming a prealar bridge. The scutum is the largest element of the notum and the
area of origin of large dorsoventral and oblique indirect flight muscles, and also of
muscles attached to the basal elements of the leg. Some smaller muscles associated
with the flight apparatus originate from its lateral regions. Laterally the scutum bears
the anterior, the postmedian and the posterior wing processes (=alar processes),
and occasionally also an anteromedian process. These small projections, usually
more or less triangular in shape, are directly connected with the wing base and interact with its axillary sclerites (see 1.3.6 Wings) (Fig. 1.3.3.1A, B). The scutal surface is
subdivided into different regions partly corresponding with the sites of origin of the
large dorsoventral muscles. The separating lines are frequently addressed as sutures
(e.g., Matsuda 1970), but are in fact ridges locally increasing the rigidity of the sclerite.
They also play a role in the context of the specific deformations resulting from alternating contractions of dorsal longitudinal and dorsoventral indirect flight muscles.
A short anterolateral scutal line separates the area bearing the anterior alar process
(occasionally also an additional anteromedian process) from the main scutal region.
In a similar way, the oblique posterolateral scutal line delimits the area bearing the
posterior alar process. The transverse transscutal