Advances in Insect Physiology, Volume 9

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Advances in

INSECT PHYSIOLOGY

VOLUME 9

This Page Intentionally Left Blank

Advances in

Insect Physiology Edited by

J. E. TREHERNE, M. J. BERRIDGE and V. B. WIGGLESWORTH Department of Zoology, The University, Cambridge, England

VOLUME 9

1972

ACADEMIC PRESS London and New York

ACADEMIC PRESS INC. (LONDON) LTD. 24/28 Oval Road, London NW1

United States Edition published by ACADEMIC PRESS INC. 1 11 Fifth Avenue New York, New York 10003

Copyright 0 1972 by ACADEMIC PRESS INC. (LONDON) LTD.

All Rights Reserved

No part of this book may be reproduced in any form by photostat, microfdm, or any other means, without written permission from the publishers Library of Congress Catalog Card Number: 63-14039 ISBN: 0-12-024209-5

PRINTED IN GREAT BRITAIN BY THE WHITEFRIARS PRESS LTD., LONDON AND TONBRIDGE

List of Contributors to Volume 9 B. BACCETTI, Institute of Zoology, University of Siena, Italy (p. 3 15) M. J. BERRIDGE, Agricultural Research Council Unit of Invertebrate Chemistry and Physiology, Department of Zoology of Cambridge, Downing Street, Cambridge, England (p. 1 ) R. G. BRIDGES, Agricultural Research Council Unit of Invertebrate Chemistry and Physiology, Department of Zoology, University of Cambridge, Downing Street, Cambridge, England (p. 5 1 ) E. M. EISENSTEIN, Department of Biophysics, 128 Chemistry Building, Michigan State University, East Lansing, Michigan 48823, U.S.A. (p. 11 1 ) P. W. MILES, School of Natural Sciences, University of Zambia, P.O. Box 2379, Lusaka, Zambia (p. 183) Y. PICHON, Institut de Neurophysiologie, C.N.R.S., Gif-mr-Yvette 92, France (P. 257) W. T. PRINCE, Agricultural Research Council Unit of Invertebrate Chemistry and Physiology, Department of Zoology, University of Cambridge, Downing Street, Cambridge, England (p. 1 ) J . E. TREHERNE, Agricultural Research Council Unit of Invertebrate Chemistry and Physiology, Department of Zoology, University of Cambridge, Downing Street, Cambridge, England (p.257)

V


Contents LIST OF CONTRIBUTORS TO VOLUME 9

. . . . . . . . . .

v

THE ROLE OF CYCLIC AMP AND CALCIUM IN HORMONE ACTION MICHAEL J . BERRIDGE AND WILLIAM T . PRINCE Introduction . . . . . . . . . . . . . . . . . . I1. The Structure and Function of Calliphora Salivary Glands . . . 111. The 5-HT-receptor Interaction . . . . . . . . . . . . IV . The Role of Cyclic AMP and Calcium as Intracellular Messengers . A. The Role of Cyclic AMP . . . . . . . . . . . . B. The Role of Calcium . . . . . . . . . . . . . . V. TheMode of Action of Cyclic AMP andcalcium . . . . . . A. The Effect of 5-HT and Cyclic AMP on Potential . . . . B. Evidence for the Independent Action of Cyclic AMP and . . . . . . . . . . . Calcium on Ion Transport C. The Time Course of the Cyclic AMP and Calcium Effects . VI . Control of Secretion-A Model of Hormone Action . . . . . VII . Comparison of 5-HT Action with that of Other Hormones . . . A. Aggregation of the Slime Mould, Dictyostelium discoideum . B. Synaptic Transmission-The Role of Cyclic AMP and Calcium inPre-andPost-synaptic Events . . . . . . . C. The Action of Epinephrine on the Heart . . . . . . . D. Excitation-Secretion Coupling . . . . . . . . . . E. Control of Metabolism . . . . . . . . . . . . . F. Transporting Epithelia-The Toad Bladder . . . . . . VIII . Conclusion . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . Notes Added in Proof . . . . . . . . . . . . . . . . . . I.

1 2 5 12 12 19 21 23 26 28 31 32 33 34 36 36 37 39 41 42 49

CHOLINE METABOLISM IN INSECTS R . G. BRIDGES I. I1.

111.

IV .

Introduction . . . . . . . . . . . . . . . . . . 5 1 Choline Metabolism in Vertebrates . . . . . . . . . . 53 Nutritional Requirements of Insects for Choline . . . . . 55 A. Choline in Insect Development . . . . . . . . . 55 B. Substitutes for Choline in the Diet of Insects . . . . 59 Water-soluble Choline Metabolites . . . . . . . . . . 63 A. Acetylcholine . . . . . . . . . . . . . . . 63 B. Phosphorylcholine . . . . . . . . . . . . . . 66 vii

viii

CONTENTS

. . Cytidinediphosphorylcholine (CDP-choline) Glycerylphosphorylcholine . . . . . . . . V . Lipid-soluble Choline Metabolites . . . . . . . . A. Phosphatidylcholine . . . . . . . . . . B. Lysophosphatidylcholine . . . . . . . . . C. Sphingomyelin . . . . . . . . . . . . VI . Enzymes Involved in Choline Metabolism . . . . . A. Cholineacetylase and Acetylcholinesterase . . . B. Enzymic Synthesis of Lipids Containing Choline C. Hydrolysis of Phosphatidylcholine . . . . . D . Oxidation of Choline . . . . . . . . . . E. Synthesis of Choline . . . . . . . . . . VII . The Metabolic Role of Choline in Insects . . . . . Acknowledgements . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . C.

D.

. . . . . . . . . . . . . . .

. . . . . . . . . . . . . . .

. . . . . . . . . . . . . .

69 70 71 71 82 83 84 84 85 87 88 89 91 100 100

LEARNING AND MEMORY IN ISOLATED INSECT GANGLIA E. M . EISENSTEIN I.

I1.

Introduction . . . . . . . . . . . . . . . . . A . The Concept of Learning . . . . . . . . . . . B. Towards a Reformulation of the Concept of Learning . C. The Use of “Model” Systems in Biology . . . . . . Behavioral Investigations . . . . . . . . . . . . . A. Intact and Headless Preparations . . . . . . . . B. Isolated Ganglion-P and R as Separate Preparations . .

.

C

IV .

V.

. .

.

112 113 115 117 118 118 128

. . . 132

Effects of Other CNS Lesions on P and R Behavior . . . Behavior of Ganglionless P and R Preparations . . . . . Discussion of P and R Leg Behavior in the Presence of Ganglionic Innervation of the Legs . . . . . . . . . G. Discussion of P and R Leg Behavior in the Absence of Ganglionic Innervation of the Legs (Ganglionless) . . . . Some Histological and Anatomical Findings Potentially Relevant to Understanding the Cellular Basis of Learning in the Cockroach . A. Introduction . . . . . . . . . . . . . . . . B. Mapping of the Cockroach Metathoracic Ganglion . . . C. Ganglion Transplantation Studies in the Cockroach . . . Electrophysiological Investigations of Learning . . . . . . A. Introduction . . . . . . . . . . . . . . . . B. Habituation Studies . . . . . . . . . . . . . C. Discussion of Habituation . . . . . . . . . . . D. Instrumental Learning Studies . . . . . . . . . . E . Classical Conditioning Studies . . . . . . . . . . F. Some Newer Electrophysiological and Analytical Approaches to Learning and Memory . . . . . . . . Molecular Approaches to Learning and Memory Storage . . . A . Introduction . . . . . . . . . . . . . . . . D. E. F.

111.

Isolated Ganglion-P and R in the Same Preparations

. . . .

136 137 140

146 149 149 149 150 150 150 150 156 157 162 164 167 167

ix

CONTENTS

B.

Assessing the Effects of Drugs on Learning and Memory in the Cockroach . . . . . . . . . . . . . . Some Speculations on the Future of the Chemical Approach C. VI . Summary . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . .

168 175 176 178 178

THE SALIVA OF HEMIPTERA PETER W . MILES I.

Introduction . . . . . . . . . . . . . . . . . . Methods of Investigation . . . . . . . . . . . . . . 111. Modes of Feeding . . . . . . . . . . . . . . . . IV. Stylet-Sheath Feeding . . . . . . . . . . . . . . . A. Sampling the Surface . . . . . . . . . . . . . Secretion of the Flange . . . . . . . . . . . . B. Formation of the Sheath . . . . . . . . . . . . C. D. Discharge of Watery Saliva . . . . . . . . . . . E. Ingestion . . . . . . . . . . . . . . . . . F. Withdrawal of the Stylets . . . . . . . . . . . . V. Lacerate-and-Flush Feeding . . . . . . . . . . . . . VI . Feeding by Carnivores . . . . . . . . . . . . . . . VII. Chemical Composition and Function of the Saliva . . . . . A . Sheath Material . . . . . . . . . . . . . . . Watery Saliva . . . . . . . . . . . . . . . . B. VIII . Phytopathogenicity . . . . . . . . . . . . . . . . IX. Salivary Glands and Ducts . . . . . . . . . . . . . A. Aphidoidea . . . . . . . . . . . . . . . . Jassomorpha . . . . . . . . . . . . . . . . B. C. Fulguromorpha . . . . . . . . . . . . . . . D. Other Auchenorrhyncha . . . . . . . . . . . . E. Heteroptera . . . . . . . . . . . . . . . . X . Origins of the Saliva . . . . . . . . . . . . . . . A . Functions of the Accessory Gland . . . . . . . . . B. Functions of the Principal Gland . . . . . . . . . C. Sources of Oxidases . . . . . . . . . . . . . D. Sources in the Homoptera . . . . . . . . . . . E. Salivary Carbohydrate and Lipid . . . . . . . . . XI . The Saliva as a Vehicle for Pathogens . . . . . . . . . . XI1. Evolution of Salivary Function in the Hemiptera: a Summary . . XI11. A Survey of Problems . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . .

I1.

183 185 191 194 194 195 196 197 200 201 202 203 205 205 208 217 225 226 229 232 233 234 236 236 237 238 239 240 241 244 247 250

THE INSECT BLOOD-BRAIN BARRIER

J . E. TREHERNE AND Y . PICHON

I. I1.

Introduction . . . . . . . . . Organization of Insect Nervous Tissues A. Extraneural Fat Body Deposits .

. . . . . . . . . . . . . . . . . . . . . . . . . . .

257 260 260

CONTENTS

X

B . The Neural Lamella . . . . . . . . . . . . . . C. The Perineurium . . . . . . . . . . . . . . . D. Glial Cells. Neurones and Extracellular Spaces . . . . . 111. The Ionic Composition of the Haemolymph and Nervous Tissues . IV . Electrical Aspects of Nervous Function . . . . . . . . . A. The Ionic Basis of Electrical Activity in Insect Nerve Cells . B. The Role of the Neural Fat Body Sheath . . . . . . . C. Neuronal Function in Intact Nervous System . . . . . D. Neuronal Function in Experimentally Treated Preparations . V . Exchanges of Radioactive Ions and Molecules . . . . . . . VI . Concluding Remarks . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . . . .

264 266 268 274 277 277 278 281 289 291 299 305 305 312

INSECT SPERM CELLS BACCIO BACCETTI I.

I1.

111.

Introduction . . . . . . . . . . . . . . . . . . 316 The Cell Surface . . . . . . . . . . . . . . . . . 317 The Acrosomal Complex . . . . . . . . . . . . . . 324 A. The Typical Triple-layered Insect Acrosomal Complex . . 324 B. The Bilayered Acrosomal Complex . . . . . . . . 326 C. The Acrosomal Complex with Only Two Outer Layers . 327 D. Monolayered Acrosomal Complex . . . . . . . . . 327 E. Total Absence of Acrosomal Complex . . . . . . . 328 The Nucleus . . . . . . . . . . . . . . . . . . 328 . . . . . . . . . . . . . . . 328 A. Nuclear Shape B . Submicroscopic Structure . . . . . . . . . . . 329 C. Chemical Characteristics . . . . . . . . . . . . 331 . . . . . . . . . . . . 331 D. Physical Characteristics The Centriolar Region . . . . . . . . . . . . . . . 332 A. The Centriole . . . . . . . . . . . . . . . . 332 B. The Centriole Adjunct . . . . . . . . . . . . . 333 C. The Initial Segment of the Axoneme . . . . . . . . 335 . . . . . . . . 336 The Axial Flagellar Filament or Axoneme A. The Microtubules . . . . . . . . . . . . . . 338 B. The Central Sheath . . . . . . . . . . . . . . 349 C. The Link-heads . . . . . . . . . . . . . . . 349 D. The Coarse Fibres . . . . . . . . . . . . . . 350 E. The Axonemal Matrix . . . . . . . . . . . . . 352 Mitochondria . . . . . . . . . . . . . . . . . . 354 A. Normal Mitochondria . . . . . . . . . . . . . 354 B . Mitochondria Transformed into Derivatives with a Crystalline Core . . . . . . . . . . . . . . . 354 C. Absence of Mitochondria . . . . . . . . . . . . 360 Accessory Ordered Flagellar Bodies . . . . . . . . . . 363 A. Structured Bodies Flanking Normal Mitochondria . . . . 363

.

IV.

V.

VI.

VII .

VIII .

xi

CONTENTS

B.

Structured Bodies Flanking the Mitochondrial Derivatives with a Crystalline Matrix . . . . . . . . . . . . C. Structured Bodies Replacing Mitochondria . . . . . . IX . Spermatozoa Possessing a Double Flagellar Apparatus or Being Devoid of it . . . . . . . . . . . . . . . . . . A. The Paired Spermatozoa . . . . . . . . . . . . B . Spermatozoa Possessing Two Axonemes . . . . . . . C. Non-flagellate Spermatozoa . . . . . . . . . . . X . Motility . . . . . . . . . . . . . . . . . . . A . Motile Mechanisms . . . . . . . . . . . . . . Metabolic Aspects of Motion . . . . . . . . . . B. The Problem of Sperm Capacitation . . . . . . . . C. XI . Spermatozoa Polymorphism and Genetics . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . .

AUTHOR INDEX SUBJECT INDEX CUMULATIVE LIST OF AUTHORS CUMULATIVE LIST OF CHAPTER TITLES

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . . . . . . . . . . . .

365 365 367 367 369 370 374 374 380 381 382 384 384 399 413 435 437


The Role of Cyclic AMP and Calcium in Hormone Action MICHAEL J. BERRIDGE AND WILLIAM T. PRINCE

A.R. C Unit of Invertebrate Chemistry and Physiology, Department of Zoology, Cambridge, England I. Introduction . . . . . . . . . . . . . . . . . . 11. The Structure and Function of Calliphora Salivary Glands . . .

The 5-HT-receptor Interaction . . . . . . . . . . . . The Role of Cyclic AMP and Calcium as Intracellular Messengers . A. The Role of Cyclic AMP . . . . . . . . . . . . B. The Role of Calcium . . . . . . . . . . . . . V. The Mode of Actionof Cyclic AMPand Calcium . . . . . . A. The Effect of 5-HT and Cyclic AMP on Potential . . . . B. Evidence for the Independent Action of Cyclic AMP and Calcium on Ion Transport . . . . . . . . . C. The Time Course of the Cyclic AMP and Calcium Effects . VI. Control of Secretion-A Model of Hormone Action . . . . . VII. Comparison of 5-HT Action with that of Other Hormones . . A. Aggregation of the Slime Mould, Dictyostelium discordeum . B. Synaptic Transmission-The Role of Cyclic AMP and Calcium in Pre- and Post-synaptic Events . . . . . . C. The Action of Epinephrine on the Heart . . . . . . D. Excitation-Secretion Coupling . . . . . . . . . . E. Control of Metabolism . . . . . . . . . . . . . F. Transporting Epithelia-The Toad Bladder . . . . . . VIII. Conclusion . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . Notes Added in Proof . . . . . . . . . . . . . . . . . . 111. IV.

.

.

1 2 5 12 12 19 21 23 26 28 31 32 33 34 36 36 37 39 41 42 49

I. INTRODUCTION

Coordination of cell activity in higher animals is achieved through various chemical messengers which are released by one cell population to influence another. Most hormones function in long range communication where the blood system carries the chemical messages to the target cells. However, communication can also occur over a short distance, such as in the synapse, where chemicals diffuse across the narrow synaptic gaps to transfer information between two AIP-2

1

2

M. J. BERRIDGE AND W. T. PRINCE

excitable cells. In both forms of communication the basic mechanism is the same-a simple chemical message is used to transfer information from one cell to another. This review attempts to summarize the progress which has been made in understanding how one particular cell system, the salivary gland of an insect, receives a chemical stimulus and translates the information into a change in activity. The salivary glands of the adult blowfly, Cullipkoru erytkrocephulu have unique structural and physiological properties which have enabled us to analyse the sequence of events which occur during cell activation by a specific hormone. Berridge and Pate1 (1 968) found that fluid secretion by isolated salivary glands was regulated by 5-hydroxytryptamine (5-HT). This simple biogenic amine may be an important secretogogue in insects because Whitehead (197 1) has evidence that 5-HT may be released from the nerves which innervate the salivary glands of the cockroach, Periplanetu americuna. In the cockroach, the increase in fluid secretion observed during nerve stimulation can be mimicked by externally applied 5-HT. Three major events have been recognized during stimulation of fluid secretion by 5-HT in the salivary glands of Cullipkoru (Berridge, 1970, 1972; Berridge and Prince, 1971, 1972a, b; Prince and Berridge, 1972; Prince et ul., 1972). Firstly, 5-HT recognizes and interacts with a specific cellular receptor. Secondly, the result of a successful 5-HT-receptor interaction is decoded into an increase in the concentration of two intracellular messengers, cyclic AMP and calcium. Thirdly, cyclic AMP and calcium are then responsible for stimulating the transport processes of the cell to produce an increase in fluid secretion. This work on Calliphoru salivary glands which has led to a simple hypothesis of the mode of action of 5-HT will form the first part of this review. In the second part, some of the general concepts which have emerged from these salivary gland studies will be discussed in relation to the mode of action of other hormones. 11. THE STRUCTURE AND FUNCTION OF CALLIPHORA

SALIVARY GLANDS

The salivary glands of the adult blowfly are paired tubules which extend down the length of the animal. That part of the gland which lies in the abdomen is uncoiled but after entering the thorax the gland forms a tangled mass which lies on either side of the

CYCLIC AMP AND CALCIUM IN HORMONE ACTION

3

proventriculus. Each tube empties into a small elongated bulb which is connected anteriorly to a cuticle-lined duct. The two salivary ducts converge and unite in the thorax to form a common duct which conveys saliva to the mouthparts. The gland has three major cell types: secretory, reabsorptive and duct cells (Oschman and Berridge, 1970). The abdominal region of the gland and most of the convolutions in the thorax consist of a single cell type (Fig. 1A). These secretory cells are characterized by extensive canaliculi formed by elaborate infoldings of the apical plasma membrane (Oschman and Berridge, 1970). These canaliculi and the free apical surface are lined with sheet-like microvilli arranged in parallel arrays. The lateral plasma membrane is relatively straight and there are septate desmosomes in the apical region. The basal plasma membrane has infoldings which are closely associated with mitochondria. These basal infoldings often come very close (less than 1 p ) to the bottom of the canaliculi. Large mitochondria with perforated cristae are scattered throughout the cytoplasm. When stimulated with 5-HT these secretory cells produce an isosmotic fluid consisting primarily of potassium and chloride (Fig. 1) (Oschman and Berridge, 1970; Berridge and Prince, unpublished). This primary secretion then passes down through a short region of the gland which is lined with a simple squamous epithelium (reabsorptive cells). Reabsorptive cells (Fig. 1B) are much less elaborate in that there are no secretory canaliculi. The apical surface has a few short microvilli resembling those of the secretory cells. The short wide basal infoldings are often closely associated with mitochondria. Various experimental procedures have revealed that this region of the gland reabsorbs potassium thus diluting the saliva (Fig. 1). This mechanism of isosmotic fluid secretion followed by hyperosmotic reabsorption to produce a dilute fluid closely resembles the mechanism of saliva production in mammalian salivary glands (Mangos e t al., 1966; Young and Schogel, 1966). That part of the gland which lies in the abdomen and is responsible for secreting the primary saliva has been used to study the mode of action of 5-HT. Secretory activity can be monitored in vitro using a technique originally devised by Ramsay ( 1954) to study isolated Malpighian tubules. When set up in control saline salivary glands secrete very slowly (0.5-1 .O nl/min) but 1 min after addition of 1 x M 5-HT the rate suddenly accelerates to more than 40 nl/min (Berridge, 1970). This high rate of secretion is maintained

4 M. J. BERRIDGE AND W. T. PRINCE

Fig. 1. The salivary gland of the blowfly. Each gland is a tubular structure divided into a secretory (A) and a reabsorptive (B) region. Schematic diagrams of the cells found in these two regions are shown. In the secretory region (A) water and chloride follow the transport of potassium resulting in the formation of an isosmotic fluid. In the reabsorptive region (B) potassium and chloride are reabsorbed so that the secreted saliva is hypotonic. bm, basement membrane; bi, basal infold; c, canaliculus;


5

for as long as 5-HT is present but returns to the resting level 1-2 min after 5-HT is withdrawn. This very rapid response and recovery implies that the 5-HT-receptor interaction is readily reversible. Since these glands are composed of a single cell type whose secretory activity can be monitored continuously, they are particularly suited for analysing the sequence of events which occur during hormone action. The first of these events is the molecular interaction between the stimulating molecule, in this case 5-HT, and its receptor. 111. THE 5-HT-RECEPTOR INTERACTION

The molecular pharmacology of 5-HT has been approached by a study of the structure-activity relationships of a wide range of analogues of 5-HT (Berridge, 1972).

I

5-HT (I) consists of an indole ring nucleus with an ethylamine chain at the 3-position and a hydroxyl group at the 5-position. The indole ring system has a high electron density with the delocalizedn electrons smeared over the two rings to form a rigid planar structure. The positions of the hydroxyl group and the 0-carbon atom are fixed by their connection to this rigid ring system. The only parts of the molecule which have any degree of freedom are the a-carbon atom and the terminal nitrogen atom of the ethylamine side chain. The activity of such compounds is usually determined by the charge distribution over the molecule. The main centre of charge resides on the ethylamine nitrogen atom which is basic and accepts a proton at physiological hydrogen ion concentrations to become positively charged (I). The hydroxyl group at the 5-position has a partial negative charge resulting from its negative inductive effect on the 71 electrons of the indole ring. These two centres of charge will be separated from each other by the hydrophobic indole ring. The activity of molecules such as 5-HT depends on two important features: (i) a high affinity for the receptor which enables them to recognize and interact with the receptor at very low concentrations and

6

M. J. BERRIDGE A N D W. T. PRINCE

(ii) once attached to the receptor they must be capable of inducing an effect often referred to as intrinsic activity. Antagonists have a high affinity for the receptor but cannot induce an effect, i.e. they have no intrinsic activity. The first experiments on 5-HT pharmacology determined which part of the molecule was responsible for its intrinsic activity (i.e. the active site on the molecule) and which part confers upon it such a high degree of specificity. A positively charged ethylamine side-chain is an important component of a wide range of biologically active compounds including histamine, y-aminobutyric acid and dopamine and is probably the active site on all these molecules. When the two major parts of the 5-HT molecule, 5-hydroxyindole and ethylamine, were tested separately they failed to activate salivary glands. When these two major parts were presented together at very high concentrations ( l oe2 M ) they still failed to stimulate secretion. The action of 5-HT thus depends on the corporate effect of both these moieties. The importance of the ethylamine chain can be demonstrated by testing a range of homologues in which the length of the chain is increased by the addition of extra carbon atoms (Fig. 2). Increasing the chain length causes a progressive increase in activity so that amylamine, which has a 5-carbon chain, stimulates the glands maximally (i.e. it has the same intrinsic activity as 5-HT). However, very high concentrations were required to stimulate secretion indicating that these simple straight chain compounds have a very low affinity for the receptor. The groups used for increasing the chain length must be hydrophobic because there is no activity if the chain terminates with a hydrophilic group as in ethanolamine (11), ethylenediamine (111) or y-aminobutyric acid (GABA) (IV). COOHCHz CH2 CH2 NH; 'NH3 CHZ CH2 NH; HOCHz CHZ NH: (11) (111) The requirement for a hydrophobic group at one end of the ethylamine chain implies that interaction of the positively charged quartenary nitrogen atom with an anionic site on the receptor requires hydrophobic interaction with the opposite end of the molecule. A progressive increase in the length of the carbon chain permits this hydrophobic interaction to develop with a resulting increase in activity (Fig. 2). That high concentrations of these compounds are needed to produce activity suggests that they have low affinities for the receptor possibly because their interaction with the hydrophobic site is unspecific.

(rv)

7


The specificity of the hydrophobic site was investigated by testing compounds where ethylamine was attached to various ring structures (Berridge, 1972). Molecules containing a benzene ring (phenethylamine) or an imidazole ring (histamine) were more active than the straight chain homologues of ethylamine, but were still very much less active then tryptamine or 5-HT which have an indole ring (Fig. 2). Thus we can conclude that the specificity of 5-HT depends on a high degree of complementarity between the indole ring and a hydrophobic site on the receptor. 5-Hydroxytryptamine

c'

N*

Tryptamine 'N

Phenethylamine C/N+ C '",

+

Histamine 0N C C." \

+ +

uc "rc c-c-c-c-N

Log concentration

I

I

I

10-s

10-4

10-3

I

+

ethylamine

10-2

(M)

Fig. 2. The dose-response curves of isolated salivary glands to 5-HT and some 5-HT-like compounds. (From Berridge, 1972.)

Another problem to consider is the role of the hydroxyl group in the 5-HT-receptor interaction. Removal of the hydroxyl group causes a thousand-fold reduction in affinity but no change in intrinsic activity because tryptamine is capable of inducing a maximal response (Fig. 2). A similar reduction in activity is observed when dimethyl tryptamine (V) is compared with the hydroxylated derivative bufotenine (VI) (Berridge, 1972). The effect of the hydroxyl group was also assessed by comparing the activity of tryptophane (VII) and 5-hydroxytryptophane (VIII). The former is totally inactive whereas the latter is capable of stimulating secretion although both the affinity and intrinsic activity are reduced. The ability of the hydroxyl group to increase the activity of the three combinations tested implies that this group increases the affinity

8


+p C-C-NH

+ /CH3

3

C-C-NH

m

Y COOH

COOH

possibly through hydrogen bonding. The position of the hydroxyl group is critical because its displacement to the 4- or 6-positions causes a thousand-fold decrease in activity similar to that observed in the absence of the hydroxyl group. These results imply that there is a hydrogen bonding site on the receptor which has a precise orientation with respect to the hydrophobic portion of the receptor which reacts with the indole ring. Having determined the contribution of the different regions of the molecule to its interaction with the receptor, it remains to establish the conformation of the ethylamine side chain as the molecule attaches itself to the receptor. As mentioned earlier, the a-carbon atom and the terminal nitrogen atom have considerable degrees of freedom and could assume various configurations as shown in Fig. 3. The &-carbon atom is free to rotate around the 0-carbon atom and can thus assume any position represented by the base of the cone I. Similarly, for most of the possible positions of the a-carbon atom, the nitrogen atom could assume a number of positions represented by the base of the second cone 11. A clue to the natural configuration of 5-HT can be obtained by testing certain molecules where this ethylamine side chain is fixed in a rigid configuration by being incorporated into additional ring systems. Preliminary unpublished experiments have shown that the lysergic acid derivative bromolysergic acid diethylamide (BOL) (IX) can stimulate secretion whereas various harmala alkaloids such as harmaline (X) are totally inactive. However, these harmala alkaloids have a high affinity for the receptor because they are potent competitors; this indicates that they have most of the attributes for a successful 5-HT-receptor interaction but once attached to the receptor they cannot induce an effect. This can be explained by a comparison of the position of the


9

HO

a

b

C

Fig. 3.

The orientation of the ethylamine sidechah of 5-HT. Rotation can occur around the p and a carbon atoms as indicated by the cones I and I1 respectively. a, b, and c show three possible conformations. Structure-activity relationship studies indicate that the active conformation is probably that in a.

nitrogen atom in the harmaline molecule with that in the active BOL molecule which suggests that the natural configuration of the ethylamine side chain is similar to that shown in Fig. 3a.

CH3

Ix

X

On the basis of these structure-activity relationships we can speculate about some of the properties of the 5-HT-receptor and the sequence of events which occur during a 5-HT-receptor interaction. The specificity of 5-HT resides in the 5-hydroxyindole ring system which suggests that the receptor must have a hydrophobic site which is precisely designed to accept the indole ring (Fig. 4).In relation to

10


I

'

Electrostatic

\ forces

I I

I I

a

b

C

Fig. 4. The proposed 5-HT-receptor interaction. The receptor consists of three major regions-a hydrophobic site, an anionic site and a hydrogen bonding site (6' ). a. The anionic site attracts the positively-charged group of 5-HTby long range electrostatic forces. b. The final approach on to the receptor depends on hydrogen bonding and van der Waal's forces. c. The indole ring occupies the hydrophobic site and allows the po8itively-charged amino group to interact with the anionic site to produce a successful 5-HT-receptor interaction.

this hydrophobic site the position of the hydrogen bonding site is critical since displacement of the hydroxyl group from the 5- to the 4-or 6-position markedly reduces activity. Situated on the opposite side of the hydrophobic site there must be an anionic site which reacts with the positively charged nitrogen atom. The results with BOL and the harmala alkaloids suggest that this anionic site has a precise location relative to the other parts of the receptor as shown in Fig. 4. Thus, we suggest that the 5-HT molecule is firmly attached to the receptor by means of three different chemical bonds: (i) an electrostatic bond with the anionic site; (ii) hydrophobic bonding (e.g. van der Waal's forces) with the hydrophobic site; (iii) hydrogen bonding involving the 5-hydroxyl group.


11

It is of some interest to speculate on the way in which these various interactions develop as 5-HT approaches and occupies the receptor. There is every reason to suppose that the event is not a haphazard collision but entails an orderly approach sequence with some degree of mutual moulding as the agonist approaches the receptor. The initial attraction and guidance towards the receptor is probably carried out by long range electrostatic forces operating between the positively charged nitrogen atom and the anionic site on the receptor (Fig. 4). Such long range forces are non-specific and a wide range of positively charged molecules will be drawn towards the anionic site. However, interaction does not occur unless there is a high degree of complementarity between the molecule and the whole receptor; in order to get closer to the receptor the approaching molecule must satisfy the more stringent requirements of the hydrophobic and hydrogen bonding sites. lndeed the hydrophobic site may protect the anionic site from the many and varied positively charged ions and molecules which will be drawn towards the negative charge on the receptor. Only 5-HT and its closely related derivatives have chemical conformations satisfying the precise requirements for combination with the hydrophobic site and the hydrogen bonding site. The hydroxyl group on 5-HT may play an additional role by ensuring that the indole ring is correctly positioned over the hydrophobic receptor site during this final approach towards the receptor. As these short-range forces pull the molecule down onto the receptor the cationic head is brought close enough to the anionic site to form an effective electrostatic bond. Probably it is this neutralization of the anionic site that initiates the hormonal effect. These 5-HT structure-activity relationships in Culliphoru salivary glands resemble those found in the isolated rat stomach strip (Vane, 1959; Offermeier and Ariens, 1966) or the heart of a mollusc (Greenberg, 1960). This basic similarity suggests that 5-HT receptors from widely different tissues possess many common features. An interesting exception to this generalization is the action of 5-HT on the Malpighian tubules of Rhodnius. Under normal conditions fluid secretion by these Malpighian tubules is stimulated by a diuretic hormone released from the fused ganglionic mass in the mesothorax (Maddrell, 1963). The action of the diuretic hormone can be simulated by 5-HT (Maddrell et al., 1971). However, structureactivity studies revealed that the receptor was very different to the 5-HT receptors described above. In particular, the presence of a hydroxyl group at the 5-position is essential for the activity of 5-HT in Rhodnius. Removal of the hydroxyl group from a series of active

12

M. J . BERRIDGE AND W. T. PRINCE

5-HT analogues converts them into powerful antagonists. In the case of these Malpighian tubules, therefore, 5-HT may be acting on the receptor normally occupied by the diuretic hormone which is thought to be a polypeptide (Maddrell, personal communication). It has been suggested that the chemical conformation of 5-HT may closely resemble the configuration of the active site on the diuretic hormone (Maddrell et al., 197 1). IV. THE ROLE OF CYCLIC AMP AND CALCIUM AS INTRACELLULAR MESSENGERS

Once 5-HT has interacted with the receptor the resulting chemical signal input is translated into intracellular messengers which are then responsible for altering cell activity. Adenosine 3',5'-monophosphate (cyclic AMP) plays a central role in mediating the action of a wide range of hormones (Robison et al., 1968). However, Rasmussen (1970) has recently suggested that calcium may be an equally important intracellular mediator of hormone action. Both these messengers appear to mediate the action of 5-HT in salivary glands. A. THE ROLE OF CYCLIC AMP

The importance of cyclic AMP in hormone action was first recognized by Sutherland and Rall(l958) who showed that it played an essential role in the hyperglycemic actions of glucagon and epinephrine on the liver. Since then this compound has been implicated in cellular control mechanisms in a wide range of organisms from bacteria to mammals (Robison et al., 1968). The concentration of cyclic AMP in most cells is determined by the balance which exists between the synthetic enzyme, adenyl cyclase, and the degradative enzyme, phosphodiesterase (Fig. 5). The level of cyclic AMP is regulated by altering the activity of adenyl cyclase which represents the site of action of most hormones as well as certain neurotransmitters (Robison et al., 1967, 1968). One action of 5-HT on salivary glands is to stimulate adenyl cyclase which increases the intracellular level of this second messenger (Berridge, 1970; Prince et al., 1972). Direct measurement of the cyclic AMP concentration showed that the control level was 0.06-0.10 p mol/gland but 2 min after the addition of 5-HT there was a 3-4-fold increase in cyclic AMP level (Fig. 6 ) . After this initial rise the concentration of cyclic AMP declines slightly to an equilibrium level which then remains constant. In the absence of calcium, the cyclic

13

CYCLIC AMP AND CALCIUM IN HORMONE ACTION Basal plasma membrane

NH2

Theophylline

5'-AMP

ATP

Fig. 5. Regulation of the internal cyclic AMP level. 5-HT stimulates the enzyme adenyl cyclase which synthesizes cyclic AMP from ATP. Cyclic AMP is hydrolysed to 5'-AMP by phosphodiesterase. The methyl xanthine theophylline can increase the intracellular concentration of cyclic AMP by inhibiting (-) phosphodiesterase.

3

I t Control

0 1

/

I

I

5

10 15 Minutes

I

I

1

20

25

Fig. 6. The effect of lO-'M 5-HT on the concentration of cyclic 3';5'-AMP in the salivary of calcium. The hatched area gland in the absence (0- - - 0 ) and presence (0-0) represents the level of cyclic AMP in unstimulated glands. (From Prince et aZ., 1972.)

14


AMP level does not increase so rapidly but it does reach and maintain a higher equilibrium level; the significance of this observation will be apparent later when the various feedback relationships between the two messengers are discussed (Section IV B). A similar effect of calcium has been observed when parathyroid hormone increases cyclic AMP levels in renal tubules (Nagata and Rasmussen, 1970). The change in intracellular cyclic AMP concentration during the action of 5-HT has also been studied by labelling the nucleotide pool with 3H-adenine. Subsequent treatment with 5-HT in the presence of the phosphodiesterase inhibitor theophylline causes a large increase in labelled cyclic AMP compared to the untreated controls. This experiment confirms that 5-HT acts to speed up the conversion of ATP to cyclic AMP as depicted in Fig. 5. The increase in cyclic AMP concentration observed on addition of 5-HT is an important link between receptor activation and the subsequent increase in fluid secretion. This direct role of cyclic AMP in cell activation is illustrated by its ability to simulate the action of 5-HT on fluid secretion (Fig. 7). The onset of the response to cyclic AMP is slightly slower than that for 5-HT. Apart from these minor temporal differences, there is a remarkable similarity in the action of these two agents. 40

-

35

-

30-

-25.E

E

-g o 1

i;

5-HT Cyclic AMP

I

.c

6$15-

I

Minutes

Fig. 7. The effect of l o - * M 5-HT, lO-’M cyclic AMP and lO-’M theophylline on the rate of fluid secretion of isolated salivary glands. All these compounds were applied at the arrow.

15


The very high concentrations of cyclic AMP required to activate secretion when applied exogenously (Figs 7 and 8) may be determined by several factors. Firstly, cell membranes may be relatively impermeable to these large nucleotides and the cell must be flooded with a very high concentration to enable enough cyclic AMP to enter the cell to raise the intracellular level sufficiently to activate secretion. Robison et al. (1965) have shown that cyclic AMP does not enter cardiac cells fast enough to achieve an effective concentration. Secondly, since phosphodiesterase is continually hydrolysing cyclic AMP, sufficient cyclic AMP must enter the cell to counteract this degradative process before stimulation of the effector system is apparent. This particular aspect can be demonstrated by studying the effect of a phosphodiesterase inhibitor on the dose-response curve for cyclic AMP (Fig. 8). After treatment with theophylline salivary glands become much more sensitive to exogenous cyclic AMP than glands without theophylline where phosphodiesterase is capable of degrading cyclic AMP maximally (Berridge, 1970). Thirdly, the natural hormone may induce an increase in other intermediates such as calcium (Section IV B) so that higher concentrations of cyclic AMP are required to overcome the lack of these other intermediates. Various cyclic AMP derivatives have been prepared in an attempt to overcome the problem of the slow rate of penetration of cyclic AMP into cells. The dibutyryl derivative of cyclic AMP (XI) introduced by Posternak et al. (1 962) has been widely employed but with very varied results. The action of the dibutyryl derivative is worth considering in some detail since it is likely to be used in many future experiments. Certain tissues such as vertebrate fat pads, salivary glands, adrenals, heart and thyroid, which are all insensitive to cyclic AMP, will respond to dibutyryl cyclic AMP (Bdolah and Schramm, 1965; Butcher et al., 1965; Imura et aZ., 1965; Pastan,

" i0-e

8

;0-7

Log concentration

1'0-6

A-5

4 0 3

I&

(MI

Fig. 8. The dose-response curves to 5-HT and cyclic AMP (c.AMP) in the presence (open circles) or absence (filled circles) of theophylline (T). (After Berridge, 1970.)

16


b -0’

b C 0 CsH-,

I 0-

XI

1966; Babad et al., 1967; Skelton et al., 1970). When applied t o Calliphora salivary glands the dibutyryl derivative stimulated secretion but the nature of the response was very different to that recorded for cyclic AMP (Berridge, 1970). The presence of substituents at the N 6 - and 2’-positions not only delay the onset of the secretory response but they also greatly prolong the recovery time after removal of the dibutyryl analogue. Considerable delays have also been observed during the action of dibutyryl cyclic AMP on the parotid gland (Babad et al., 1967) isolated fat cells (Perry and Hales, 1970) and heart (Skelton et al., 1970). There is some indication that the potency of cyclic AMP depends on having a free hydroxyl group at the 2’-position on the ribose ring (Henion et al., 1967). Consequently, the slow response of salivary glands and other tissues to dibutyryl cyclic AMP may depend on the time required for intracellular esterases t o remove the butyryl substituent from the 2‘-position before this derivative can become active. The slow recovery after removing dibutyryl cyclic AMP from salivary glands (Berridge, 1970) may be explained by the observation that this compound is less susceptible than cyclic AMP t o degradation by phosphodiesterase (Posternak et al., 1962). All these factors must be taken into consideration when attempting t o interpret results obtained with these substituted derivatives. The specificity of cyclic AMP has been tested by comparing its response with a number of related cyclic nucleotides (Fig. 9). Cyclic nucleotides consist of three major regions: (i) a purine or pyrimidine base (R in Fig. 9); (ii) the sugar ribose which is linked to the base by a glycosidic bond; (iii) a phosphate group which forms a ring through its connection to the 3’- and 5’-carbon atoms of the ribose sugar (see Fig. 5).


17

The structure-activity studies reveal that the cyclic AMP receptor can accommodate certain modification of the base region (Fig. 9). Indeed the tubercidin and uridine cyclic phosphates were more active than cyclic AMP. Drummond and Powell (1 970) have also observed that cyclic tubercidin 3',5'-monophosphate is more active than cyclic AMP in stimulating phosphorylase b kinase from skeletal muscle. In salivary glands cyclic inosine monophosphate is less active than cyclic AMP and also shows a reduction in intrinsic activity in that it cannot induce a maximal effect (Fig. 9). Cyclic guanosine monophosphate is totally inactive even though this compound has been identified in various vertebrate tissues (Hardman et aL, 1971). All compounds tested which contained modifications of the ribose or phosphate rings were totally inactive. Some of the most interesting compounds tested were 5'-adenosine monophosphate (see Fig. 5), desoxyadenosine 3',5'-monophosphate (XII), adenine 9-/3-D-xylofuranosyl 3',5'-monophosphate (XIII) and adenosine 3',5'-cyclic phosphorothioate (XIV). In 5' AMP the phosphate ring has been opened (the 5'derivatives of various other cyclic nucleotides were also totally inactive, e.g. 5'UMP, 5'IMP, 5'GMP). In XI1 the 2'-hydroxyl group is missing. The 3'-hydroxyl group in XI11 is positioned above the plane of the ring which allows the formation of an unstrained phosphate R

10. Log concentration (M)

Fig. 9. The dose-response curves of salivmy glands produced by different cyclic 3'3'-nucleotide monophosphates. c.TMP-cyclic tubercidin monophosphate. c.UMP-cyclic uridine monophosphate. c.IMP-cyclic inosine monophosphate. c.GMP-cyclic guanosine monophosphate.The structure of the different bases (R) is shown.

18


diester linkage in contrast to the bond in cyclic AMP which is very strained (Drummond and Powell, 1970). Activity is also lost if the double bond oxygen is replaced with the much bulkier sulphur atom (XIV). The latter compound is of particular interest because it is the only nucleotide tested which appeared to compete with cyclic AMP. There have been few reports of cyclic AMP competitors. Kulka Adenine

Adenjne

Adenine

and Sternlicht (1 968) have shown that adenosine 3'-monophosphate may inhibit the ability of cyclic AMP to stimulate enzyme secretion in mouse pancreas. Adenosine 3'-monophosphate had no effect on salivary glands. Further evidence to implicate cyclic AMP in the control of secretion has come from studies with the phosphodiesterase inhibitor theophylline (Berridge, 1970). By reducing the breakdown of cyclic AMP within the cell, theophylline can lead to higher than normal intracellular levels of cyclic AMP which in turn lead to several predictable consequences. Firstly, theophylline can stimulate secretion directly but the onset of the response takes much longer than that for either 5-HT or cyclic AMP (Fig. 7). Evidence will be presented later (page 27) t o show that the long delay represents the time required for the unstimulated adenyl cyclase to raise the internal cyclic AMP concentration high enough to stimulate secretion. The ability of theophylline to potentiate the action of cyclic AMP has already been described (Fig. 8 ) and this potentiation can extend to the action of the primary stimulus 5-HT. If the internal destruction of cyclic AMP is decreased, it becomes much easier for adenyl cyclase to raise the cyclic AMP concentration high enough to stimulate secretion. In other words, adenyl cyclase need not be activated maximally which in turn means fewer 5-HT-receptor interactions and therefore much lower concentrations of 5-HT are required. Figure 8 illustrates the marked potentiation which can be obtained with theophylline. The dose-response curve for 5-HT is shifted to the left. Thus a simple chemical signal, 5-HT, arriving at the surface is


19

translated into an intracellular secondary messenger in the form of cyclic AMP through the activation of the enzyme adenyl cyclase. B. THE ROLE OF CALCIUM

The actions of both 5-HT and cyclic AMP are critically dependent upon calcium. In the absence of calcium the onset of the secretory response of salivary glands to 5-HT and cyclic AMP is slower than normal (cf. Figs 7 and 10). This is particularly true for the action of cyclic AMP which showed a 2-min delay before the onset of secretion in a calcium-free medium. In both cases, a brief stimulation of secretion was followed by a decline to very low rates of secretion. In the case of the 5-HT-stimulated glands, this inability to secrete in a calcium-free environment is not caused by any impairment of adenyl cyclase activity since the cyclic AMP level under such conditions is higher than in the presence of calcium (Fig. 6). The limiting factor is calcium availability as shown by the dramatic recoveries evident when this divalent cation is returned to the bathing medium (Fig. 10). Flux studies with radiocalcium show that 5-HT, but not cyclic AMP, stimulates 45Ca influx into isolated salivary glands (Prince et a l , 1972). The efflux of 45Ca from prelabelled cells, however, was stimulated by both 5-HT and cyclic AMP. The ability of both agents to influence calcium efflux points to a complicated feedback

0

10

20

30

40

50

60

Minutes

Fig. 10. The effect of l o - * M 5-HT(0-0) and lo-’ M cyclic AMP (0- - - 0 ) applied at the first arrow,on the rate of fluid secretion in the absence of calcium. 2.0 mM calcium was added to the bathing medium at the second arrow. (After Prince et oZ., 1972.)

20

M . J. BERRIDGE AND W. T. PRINCE

relationship operating between these two second messengers. There is some evidence that cyclic AMP can affect intracellular calcium levels by influencing the intracellular pools of calcium contained in the mitochondria or sarcoplasmic reticulum (Rasmussen, 1970; Epstein et al.. 1971). The ability of 5-HT and exogenous cyclic AMP to increase calcium efflux from salivary glands could be explained if increasing the intracellular concentration of cyclic AMP leads t o greater release of calcium from some intracellular pool. This ability of one intracellular messenger to influence the concentration of the other messenger raises an important question concerning the role of various feedback relationships during hormone action. These feedback mechanisms are poorly understood but they are sufficiently important to warrant some discussion at this early stage because they may further our understanding of intracellular calcium homeostasis. The free calcium which enters the cytoplasm can originate from both the external bathing medium and from intracellular pools such as the mitochondria. One action of 5-HT is to increase calcium influx from the external medium and there is circumstantial evidence from the efflux studies that cyclic AMP exerts a positive feedback control on the release of calcium from some intracellular pool (Fig. 11). The slow onset of secretion Basal plasma membrane

Ca2

+

5-HT

/“O AMP

u lntracellular calcium pool

Fig. 11. The interrelationships of calcium and cyclic AMP in the salivary gland. The cytoplasmic calcium is derived from two sources: the external medium and an intracellular pool, probably mitochondria. Cyclic AMP may release calcium from this pool (+) and the calcium may inhibit adenyl cyclase (-) thus controlling cyclic AMP levels. The entry of calcium and the production of cyclic AMP are both stimulated by 5-HT.


21

induced by 5-HT and cyclic AMP in a calcium-free medium (Fig. 10) may depend on this proposed action of cyclic AMP on calcium release from the intracellular pool. As this calcium enters the cytoplasm, secretion is possible and accounts for the temporary burst of fluid secretion (Fig. 10). However, as this calcium is lost to the external calcium-free environment and is not replaced, secretion fails as the intracellular calcium reserves are depleted. Calcium, in turn, might be able to regulate cyclic AMP levels through a negative feedback control on adenyl cyclase activity as suggested by Chase et al. (1 969). This divalent cation inhibits adenyl cyclase activity in broken cell preparations of heart (Drummond and Duncan, 1970), kidney (Streeto, 1969), and foetal rat calvaria (Chase et al., 1969). The increased intracellular levels of cyclic AMP found in salivary glands incubated in a calcium-free medium (Fig. 6) is certainly consistent with such a feedback relationship. A similar mechanism could account for the higher than normal levels of cyclic AMP observed in calcium-free solutions during the action of epinephrine on the heart (Namm et al., 1968) and parathyroid hormone on the kidney (Nagata and Rasmussen, 1970). It is not claimed that the internal control mechanism in salivary glands can be described in terms as simple as the feedback relationships depicted in Fig. 1 1. There may be more feedback loops and more complex modes of interaction of the various processes which regulate the concentrations of cyclic AMP and calcium. The important point to establish at this stage is that such feedback relationships may exist and may play an important role in integrating hormone action within the cell. These various feedback mechanisms may be considered as sophisticated devices to ensure that the rapid changes in cell activity produced by hormonal stimuli are orderly and regulated. These devices may also prevent the second messenger concentrations from exceeding that required for a maximal activation of the cell. An important consequence of keeping such a check on the concentration of the second messengers is that it enables the cells t o recover rapidly on withdrawal of the hormonal stimulus. V. THE MODE OF ACTION OF CYCLIC AMP AND CALCIUM

The way in which the two intracellular messengers, cyclic AMP and calcium, react with the transport mechanisms responsible for

22

M. J . BERRIDGE AND W. T. PRINCE

secreting fluid has been investigated using electrophysiological techniques. The osmotic gradients necessary to promote a passive flow of water across salivary glands are generated by the transport of potassium and chloride (Oschman and Berridge, 1970; Prince and Berridge, 1972, unpublished observations). Ion transport across an epithelium usually generates an electrical potential which can be used to analyse the nature of the underlying ionic events. We have used this approach to try and uncover the site and mode of action of cyclic AMP and calcium. The sucrose-gap technique used by neurophysiologists has been adapted to study the electrical responses of isolated salivary glands (Berridge and Prince, 1971, 1972a, b; Prince and Berridge, 1972). The salivary gland (SG) is ligated at both ends with fine silk threads (SL) and transferred to a perspex chamber consisting of three parallel baths (Fig. 12). The gland which lies in a groove connecting all three baths is held in position by inserting the ligatures into Vaseline on the two outer walls. The closed end of the gland lies in the perfusion bath (PB) which is constantly perfused with saline (arrows). The open end of the gland lies in the saliva bath (SB) which also contains saline so that the lumen of the gland is in electrical contact with this bath. These two outer baths are also insulated from each other by liquid paraffin (LP) contained in the middle bath. The transepithelial PB

Fig. 12. Diagram of the perspex perfusion chamber used to record the electric potentials of isolated salivary glands. (From Berridge and Prince, 1972a.) See text for further details.


23

potential can be measured by connecting each outer bath to an electrometer through agar bridges (AB) and calomel electrodes. Intracellular potentials can be recorded by inserting a microelectrode into gland cells lying in the perfusion bath (Prince and Berridge, 1972). Agar blocks (A) support the gland during penetration with electrodes and also increase mixing in the perfusion bath. A. THE EFFECT OF 5-HT AND CYCLIC AMP ON POTENTIAL

The transepithelial potential of resting glands was approximately 4mV with the lumen positive to the bathing medium but a few seconds after treatment with 5-HT this potential changed to about 16 mV negative of the resting value and remained at this level as long as 5-HT was perfused over the gland (Fig. 13). After removal of 5-HT

"b

Fig. 13. The electrical response of an isolated salivary gland to 5-HT (lo-' M). The positions of the recording and reference electrodes for each trace are shown on the right; the hatched area represents the liquid paraffin gap which insulates the perfusion bath, containing the closed end of the gland, from the saliva bath. a. The transepithelial potential (VT ) recorded by means of calomel electrodes connected to the two outer baths. b. The potential across the basal surface (V,,), recorded by means of a microelectrode with the perfusion bath calomel electrode as a reference (note that the recording across this surface is greatly amplified). c. The potential across the apical surface (V,) recorded by using the calomel electrode in the saliva bath as a reference. The potential change across this surface is very similar to the change in transepithelial potential. (From Berridge and Prince, 1972b.)

24


the potential returned to its resting level. The transepithelial potential is the sum of the potential changes occurring across the basal and apical surfaces of the salivary gland cells. Intracellular microelectrode recordings revealed that most of the transepithelial potential can be accounted for by a large depolarization of the apical plasma membrane (Fig. I3c). The small hyperpolarization (2-4 mV) across the basal plasma membranes developed much more slowly than that across the apical surface and may simply reflect changes in the concentration of intracellular ions which develop during the increased ion flux induced by 5-HT. The effect of cyclic AMP on potential (Fig. 14) was very different from that just described for 5-HT even though these two agents have such similar stimulatory effects on secretion (Fig. 7). Instead of going negative as with 5-HT, the transepithelial potential went even more positive after treatment of salivary glands with 1W 2M cyclic AMP (Fig. 14). Analysis of this response again revealed that the potential change occurs predominantly across the apical surface. Cyclic AMP caused the apical membrane to hyperpolarize. The small change which occurred across the basal plasma membrane was similar to that recorded on addition of 5-HT.

U

Cyclic AMP

lmin

Fig. 14. The electrical response of an isolated salivary gland to cyclic AMP (10- '11). The position of the recording and reference electrodes are shown on the right as described in Fig. 13. a. The transepithelial potential (V,) shows an increase in lumen positivity after treatment with cyclic AMP. b. There is a small hyperpolarization across the basal surface (V,). c.Most of the transepithelial potential can be accounted for by a large hyperpolarization of the apical surface (VJ. (From Berridge and Prince, 1972b.)


25

These potential responses to 5-HT and cyclic AMP can be summarized in the form of potential profiles shown in Fig. 15. 5-HT causes the transepithelial potential to go negative through a large depolarization of the apical plasma membrane (Fig. 15b). In contrast, cyclic AMP induces an increase in positivity by hyperpolarizing this apical surface (Fig. 15c). These observations apparently cast some doubt on the proposed role of cyclic AMP as a second messenger because if it was an intermediary it should exactly *30-

'20 +lomV

0-10

-

-20 -30 -40 -50 -60Resting

5-HT

Cyclic AMP

Fig. 15. A summary of the average potentials recorded across the basal (B) and apical (A) surfaces of the salivary gland cells in the resting state and during stimulation with either 5-HT or cyclic AMP. The opposite effects of 5-HT and cyclic AMP on the transepithelial potential (hatched area) are restricted to changes across the apical surface. 5-HTdepolarizes whereas cyclic AMP hyperpolarizes this membrane. (From Berridge and Prince, 1972b.)

duplicate the effects of the first messenger 5-HT on the transepithelial potential. However, this dilemma can be resolved if 5-HT has two actions only one of which is mediated by cyclic AMP. The electrical recordings indicate that cyclic AMP stimulates cation transport to account for the large increase in positivity seen when this compound acts in the absence of 5-HT. The increase in negativity observed with 5-HT suggests that there is stimulation of anion transport which is independent of cyclic AMP. Subsequent studies (e.g. Prince et al, 1972) which will be described later, have shown that this action of 5-HT on anion transport is mediated by calcium. Thus the two intracellular mediators of 5-HT action have very different effects on the ionic mechanisms responsible for secretion. The electrical measurements suggest that potassium transport is driven by an electrogenic pump located on the apical plasma membrane whereas chloride movement is mainly passive and is linked t o active cation transport. The potential generated across the gland depends both on the activity of the electrogenic pump and also on the ease with which

26


chloride ions can enter the lumen to counteract this flow of cations (Prince and Berridge, 1972). Cyclic AMP appears to regulate the active cation pump whereas the intracellular calcium concentration determines either the permeability to anions or the degree of linkage of anions to cation transport. Evidence for this hypothesis has been obtained from a number of experiments designed to discriminate between the two different actions of 5-HT. The interpretation of such experiments must take into account the complex feedback relationships which may exist between cyclic AMP and calcium mentioned earlier (Fig. 11). Since the regulation of these two messengers is intimately related there are a limited number of procedures for obtaining an increase in the concentration of one without a simultaneous rise in the other. Most of the experiments are concerned with studying the effects of increased levels of cyclic AMP in the absence of calcium. B. EVIDENCE FOR THE INDEPENDENT ACTION OF CYCLIC AMP AND CALCIUM ON ION TRANSPORT

The simplest way of increasing the internal concentration of cyclic AMP without the mediation of 5-HT is to add this nucleotide to the bathing medium. The resulting increase in positive potential described earlier (Fig. 14) is consistent with the idea that cyclic AMP stimulates a cation pump and consequently increases the apical membrane potential. Since cyclic AMP mimics the action of 5-HT on secretion (Fig. 7), chloride movement must be high enough to support the high rate of cation transport necessary for a maximal rate of secretion during the action of exogenous cyclic AMP. It is conceivable that the ability of cyclic AMP to release calcium from an intracellular pool (as postulated in Fig. 1 1 ) keeps the internal calcium concentration just above the critical level for solute transport but not high enough to produce depolarization of the apical membrane. However, when salivary glands were treated with 5-HT or cyclic AMP in a calcium-free medium the internal calcium levels, and hence anion movement, were apparently limiting because the rate of secretion was very low (Fig. 10). In calcium-free media the internal cyclic AMP concentration is high, as measured directly in the presence of 5-HT (Fig. 6). In a calcium-free medium, 5-HT and cyclic AMP produce large positive potentials which are again consistent with an action of cyclic AMP on cation transport in the absence of the calcium-dependent increase in chloride movement

27


(Prince et al., 1972; Berridge and Prince, 1972b). When calcium is returned to the medium bathing a gland stimulated by 5-HT in a calcium-free medium, the transepithelial potential immediately returns to the negative value characteristic of the action of 5-HT in normal saline (Berridge and Prince, 1972b). Another way of studying the action of cyclic AMP in the absence of the 5-HT-induced increase in calcium influx is to use the phosphodiesterase inhibitor theophylline. As is discussed on p. 18, theophylline can stimulate secretion by inhibiting the intracellular breakdown of the cyclic AMP generated by the unstimulated adenyl cyclase. As one might expect, there is an increase in positive potential which has a very slow onset like that observed for the onset of secretion (Fig. 16) (Berridge and Prince, 1971). The slow time

t20L I

0

1

I

I

1

I

II)

I

I

I

I

1

20

I

I

I

I

timelmin

Fig. 16. The effect of l O - * M theophylline (horizontal bar) on rate of fluid secretion (nl/min) and the transepithelial potential (mV)of isolated salivary glands. (From Berridge and Prince, 1971.)

course for the onset of the secretory and potential response is probably indicative of the slow build up in the concentration of intracellular cyclic AMP which relies on the activity of the unstimulated adenyl cyclase. If this is the case, it should be possible to induce the action of theophylline much more rapidly by increasing the activity of adenyl cyclase. If at the beginning of the theophylline treatment, the turnover of adenyl cyclase is briefly enhanced by a 6 s treatment with 5-HT, the positive potential develops more rapidly (Berfidge and Prince, 1972a). This experiment clearly illustrates that the delayed secretory and potential responses

28


observed during theophylline action cannot be accounted for by any delay in the development of phosphodiesterase inhibition. Indeed the inhibition develops very fast and the long delays depend solely on the time required for the unstimulated adenyl cyclase to raise the cyclic AMP concentration to the level necessary to stimulate the secretory and potential responses of the cell. It is also possible to distinguish between the different actions of cyclic AMP and calcium by preventing the expression of their effects by altering the ionic composition of the bathing medium. If salivary glands are stimulated with 5-HT in a saline where chloride has been

1

0

1

1

1

1

5

1

1

1

1

~

10

1

timelmin

Fig. 17. The transepithelial potential response of isolated salivary glands to lO-'M 5-HT (horizontal bar) when all the chloride in the saline was replaced with isethionate. (From Berridge and Prince, 197 1 .)

replaced with the large impermeant anion isethionate, the normal increase in negativity is not seen and the lumen becomes strongly positive (Fig. 17). Under these conditions 5-HT will function normally to increase the intracellular concentrations of both cyclic AMP and calcium. The former will stimulate cation transport but the action of calcium on anion movement is not expressed because isethionate cannot permeate the cell, the net result is a large increase in positivity (Berridge and Prince, 197 1, 1972a). C. THE TIME COURSE OF THE CYCLIC AMP A N D CALCIUM EFFECTS

A careful analysis of the electrical events which occur during stimulation with 5-HT in different conditions has revealed that the time course of the actions of the two messengers are very different. The increase in negativity, which is thought to reflect an increase in anion movement caused by a rise in the internal calcium concentration, develops very rapidly with a half time of about 5 s. The time course for the increase in the internal cyclic AMP


29

concentration is more difficult to determine but the isethionate experiments just described provide an indirect measure of the build up of cyclic AMP. The method depends on the assumption that during the action of 5-HT in an isethionate saline, the action of cyclic AMP can be unmasked since the movement of anions is prevented. In an isethionate saline, the increase in positivity has a half time of 30 s which is much longer than the increase in negativity induced by 5-HT in a normal chloride-containing saline (half time of 5 s). Thus, these measurements indicate that during the action of 5-HT the build up of calcium and cyclic AMP within the cell occurs at very different rates. The action of 5-HT on calcium influx leads to a very rapid increase in internal calcium concentration which in turn stimulates chloride movement with a rapid increase in negativity. The increase in cyclic AMP concentration, which depends on the action of 5-HT on adenyl cyclase, occurs much more slowly. The slow increase in cyclic AMP concentration may account for the delay in the onset of fluid secretion after administration of 5-HT. Although calcium will flood into the cell very rapidly and activate chloride movement, fluid secretion will not begin until active cation transport is stimulated by cyclic AMP. The half time for the onset of secretion is 35 s (Fig. 7) which is very close to the indirect estimate from potential measurements of 3 0 s for the build up of internal cyclic AMP concentration (Berridge and Prince, 1972a). The different time course for the actions of the two intracellular messengers provides another way of distinguishing their separate effects under normal conditions. If salivary glands are treated with 5-HT for very short periods, the normal increase in negativity is always followed by a marked increase in positivity. This “positive undershoot” gradually returns to the normal resting potential. The hypothetical curves in Fig. 18 attempt to describe this biphasic response in terms of the postulated differences in the time course for the actions of cyclic AMP and calcium. During a brief treatment with 5-HT, calcium will rapidly flood into the cell to cause an immediate increase in negativity through its action on chloride movement. At the same time 5-HT will also begin to activate adenyl cyclase to increase the intracellular concentration of cyclic AMP. At about the time the cyclic AMP level begins to rise, 5-HT begins to leave the bath causing an immediate decline in calcium influx and a subsequent return of the internal calcium concentration to unstimulated levels. The cyclic AMP system apparently does not respond so rapidly to the removal of 5-HT. The marked positive undershoot can

30

M. J. BERRIDGE AND W. T. PRINCE a

-20-

-10

-

0-

> E

+lo-

+20

-

t30

-

b

I

0

~

1

I

2

I

1

3 4 5 Minutes

'

6

'

7

'

'

8

Fig. 18. a. 'Ihe response of an isolated salivary gland to treatment with 1 x lO-'M 5-HT for 15 s (horizontal bar). b. Hypothetical curves illustrating the proposed changes in internal and cyclic AMP (---) concentrations during the response shown in a. The calcium (-) stippling represents the period when cyclic AMP acts in the absence of calcium which may account for the positive phase of the biphasic response in a.

be interpreted on the basis that cyclic AMP continues to activate cation transport for a short period after the action of 5-HT on anion movement has waned. The stippling represents the period when the internal cyclic AMP concentration is elevated in the absence of calcium (Fig. 18b). The steepness of the rising phase of this parameter may account for the sudden positive potential plunge observed during the biphasic response to short treatments of 5-HT (Fig. 18a). The recovery of the potential possibly indicates the destruction of cyclic AMP by phosphodiesterase. Some proof for the involvement of cyclic AMP in the development of the positive


31

undershoot has come from studies using theophylline which can stabilize the potential at the bottom of the positive undershoot and prevent the gradual recovery to an unstimulated potential (Berridge and Prince, 1972a). VI. CONTROL OF SECRETION-A MODEL OF HORMONE ACTION

Since thidiscovery of cyclic AMP by Sutherland and Rall(1958) this compound has been implicated in the control of a wide range of cellular processes. The original hypothesis, which has gained wide acceptance, that the actions of most hormones are mediated solely by cyclic AMP was clearly an oversimplification and Robison et al. (1968) have suggested that there may be other second messengers. Recently it has been postulated that calcium may have an equally important role in mediating hormone action (Rasmussen, 1970). This idea is particularly appealing because intracellular ions are known to play an important role in regulating many cellular functions (Bygrave, 1967). These studies on salivary glands suggest a model of hormone action which may be applicable to other hormones. The primary hormonal message arriving at the salivary gland cell is carried in the chemical configuration of the 5-HT molecule. This information is decoded into secondary messages in the form of cyclic AMP and calcium through a specific interaction with a cellular receptor (Fig. 19). The most important aspect of the 5-HT-receptor interaction appears to be the formation of an electrostatic bond between the positively charged quartenary nitrogen atom at the end of the ethylamine side chain and an anionic site on the receptor. The specificity of the response depends on precise hydrophobic and hydrogen bonding involving the indole ring and its hydroxyl group with corresponding sites on the receptor. The perturbation induced in the receptor as a result of this 5-HT interaction is then translated into an increase in the concentration of two intracellular messengers. The first event appears to be an increase in intracellular calcium concentration which may arise through a change in the calcium permeability of the basal plasma membrane. The simultaneous activation of the enzyme adenyl cyclase leads to a slightly slower increase in the internal cyclic AMP concentration. All further actions of 5-HT on cell activation are mediated by these two intracellular messengers. Cyclic AMP acts by stimulating cation transport whereas calcium functions by increasing the flow of anions across the apical plasma membrane. More information on the ionic basis of secretion

32

M . J. BERRIDGE AND W. T. PRINCE Apical membrane

Basal membrane

wir

I

I '

.- . -. ,

-.

Fig. 19. A model for hormone action. 5-HT acting on the receptor (stippling) at the basal membrane stimulates two processes: the entry of calcium and the production of cyclic AMP. The increase in the level of cyclic AMP stimulates the pumping of potassium across the apical membrane. Calcium controls the movement of chloride. Water follows the movement of potassium and chloride resulting in secretion. For clarity the details of cyclic AMP and calcium regulation (shown in Figs 5 and I 1 )are omitted.

is required before the actions of cyclic AMP and calcium can be described in greater detail. VII. COMPARISON OF 5-HT ACTION WITH THAT OF OTHER HORMONES

Animal hormones can be divided into two main groups. One group is concerned with regulating growth and development and is usually thought to act via the genome. Juvenile hormone and ecdysone are two such insect hormones which determine the long-term structure and function of cell populations. The mode of action of such hormones have been reviewed by Wigglesworth ( 1964, 1970) and Highnam and Hill (1969). It has been suggested that cyclic AMP may be involved with the action of ecdysterone (Leenders et al., 1970). The second group of hormones is more concerned with regulating the day-today activity of cells at each developmental stage during the life of the animal. For example, insect hormones have been discovered which regulate the transient colour changes observed in mantids (reviewed in Wigglesworth, 1970), the contraction of heart and visceral muscles (Davey, 1964), the conversion of glycogen to trehalose by fat body (Steele, 1963) and the activity of the excretory


33

system (reviewed by Highnam and Hill, 1969). The Malpighian tubules of Rhodnius are regulated by a diuretic hormone released from the fused ganglionic mass in the mesothorax (Maddrell, 1963). The action of this hormone may have many similarities to that of 5-HT on Culliphoru salivary glands. One action of the diuretic hormone is to increase the permeability of the Malpighian tubules to chloride ions and this accounts for a sudden increase in negativity which precedes the onset of secretion by several minutes (Maddrell, 1972). The subsequent activation of cation and anion pumps then induces a biphasic potential response associated with the beginning of fluid secretion. The latter response may depend on cyclic AMP which is thought to mediate the action of the diuretic hormone of Rhodnius (Maddrell et ul., 1971). Apart from these studies on the diuretic hormones of Rhodnius there is very little information on the mode of action of this second group of hormones in insects. The present study on the control of fluid secretion by salivary glands may provide a basic model for hormones concerned with regulating the short-term activity of insect cells. Due to our lack of knowledge about the mode of action of insect hormones, the general significance of this model for the control of secretion by salivary glands must be assessed by comparing the mode of action of 5-HT (summarized in Fig. 19) with that of hormones found in other animal groups. The following general features of the model outlined in Fig. 19 will be considered: (i) Interaction of the hormone with the receptor causes a change in membrane permeability to calcium, and possibly to other ions as well, in addition to adenyl cyclase activation. (ii) The increase in intracellular concentration of cyclic AMP and calcium mediate all further actions of the hormone. (iii) The intracellular concentrations of calcium and cyclic AMP are controlled by various internal feedback mechanisms. The following examples are mainly taken from vertebrate systems. In some examples the action of the hormone is not fully understood but they all display certain features seen in the action of 5-HT on salivary glands. In every case cyclic AMP and calcium are implicated in the control of cellular activity. A. AGGREGATION OF THE SLIME MOULD, DICTYOSTELIUM DISCOIDEUM

This slime mould can exist in two forms, a motile single-cell amoeba or aggregates of hundreds of individuals differentiated into a AIP-3

34


spore. The ability of cyclic AMP to induce the motile individuals to aggregate (Konijn et al., 1967) requires the presence of calcium (Mason et al., 197 1). Chi and Francis (1 97 1) have shown that cyclic AMP stimulates calcium efflux from prelabelled amoebae suggesting a mobilization of intracellular stores as noted in salivary glands (p. 20). Thus an important relationship between cyclic AMP and calcium can be recognized even in this very primitive system. B. SYNAPTIC TRANSMISSION-THE ROLE OF CYCLIC AMP AND CALCIUM IN PRE- AND POST-SYNAPTIC EVENTS

Some of the most active preparations of adenyl cyclase have been obtained from nervous tissue (Sutherland et al., 1962; Weiss and Costa, 1968). Both electrical stimulation and treatment with catecholamines promotes an accumulation of cyclic AMP in brain slices (Kakiuchi and Rall, 1968a, b; Kakiuchi et al., 1969). There is no evidence that cyclic AMP plays a direct role in synaptic events but it may function in neural integration by modulating pre- and post-synaptic events. Although it is dangerous to generalize when so little is known, there are clear indications that cyclic AMP may facilitate pre-synaptic events but inhibit post-synaptic activity. Further, cyclic AMP also appears t o mediate most of the synaptic events which are controlled by catecholamines.

Pre-sy nap tic Activity The primary role of calcium in regulating the release of a wide range of cellular secretions is well established (Simpson, 1968; Smith, 1971). A role for cyclic AMP in the release process of cells including synaptic vesicles has also been recognized. The control of hormone and enzyme release will be described in greater detail later. Direct evidence for an involvement of cyclic AMP in pre-synaptic events has come from studies on neuromuscular transmission. Catecholamines can facilitate neuromuscular transmission by promoting the release of acetylcholine from motor neurones (KrnjeviC and Miledi, 1958; Jenkinson et al., 1968). This action of catecholamines can be simulated by both dibutyryl cyclic AMP and theophylline (Breckenridge et al., 1967; Goldberg and Singer, 1969) which can increase the frequency of the miniature end-plate potentials (m.e.p.p.’s). The exact mode of action of cyclic AMP is unclear, but Singer and Goldberg (1 970) suggest that it may function by altering calcium levels. Since external calcium concentration has


35

an important effect on the frequency of m.e.p.p.’s (Hubbard et al., 1968), cyclic AMP may modulate synaptic transmission by adjusting the intracellular level of calcium. Whether or not the present observations on neuromuscular transmission can be extended to the release of transmitters in the central nervous system is unknown. However, the observation that the very high activities of adenyl cyclase and phosphodiesterase are located in the synaptosomal fraction (DeRobertis et al., 1967) is very suggestive. Post-synaptic Activity The action of catecholamines in post-synaptic events in both the central and peripheral nervous system may also depend on cyclic AMP. Cyclic AMP mediates the inhibitory action of norepinephrine on the spontaneous discharge rate of cerebellar Purkinje cells (Siggins et al., 1969, 197 la). Intracellular recordings reveal that inhibition results from a large hyperpolarization achieved through an increase in membrane resistance (Siggins e t al., 197 1b). Dopamine has a similar inhibitory action in the peripheral nervous system (McAfee et al., 1971). Dopamine released from interneurones stimulates adenyl cyclase on the post-ganglionic neurone and the resulting increase in cyclic AMP leads to a slow wave of hyperpolarization. Cyclic AMP also plays an important role in the function of smooth muscle. As in other types of muscle, calcium is the agent responsible for the coupling between excitation and contraction. The contractile state of the muscle is thought to be related to the free calcium level in the cytoplasm; it is this level which is altered by excitatory and inhibitory transmitters. In most smooth muscles the action potentials induced by the excitatory transmitters are responsible for triggering contraction. During each action potential, calcium which is the charge carrier (Brading e t al., 1969; Bulbring and Tomita, 1970) enters the cytoplasm and induces a brief mechanical response (phasic response). There must be a rapid train of impulses in order to produce a full contracture. Cyclic AMP is apparently not involved in the development of such contractures but appears to be associated more with the relaxant effect of catecholamines. The possibility that cyclic AMP relaxes muscles by reducing the calcium level is suggested from studies on the uterus where the ability of calcium to induce contractions can be blocked by cyclic AMP (Mitznegg e t al., 1970). 0-Adrenergic agents induce an increase in the cyclic AMP content of smooth muscle from the intestine (Bueding et al., 1966) and uterus (Dobbs and Robison, 1968). This increase in cyclic AMP hyper-

36

M. J. BERRIDGE AND W.

T.

PRINCE

polarizes the membrane possibly through the activation of a sodium and/or calcium pump (Somlyo et al., 1970). Both cyclic AMP and calcium are thus intimately connected with the ability of catecholamines to modulate the activity of excitable cells. C. THE ACTION OF EPINEPHRINE ON THE HEART

Myocardial contractility is modulated by epinephrine which increases both the force of contraction (inotropic effect) and activates phosphorylase to accelerate glycogenolysis. After the administration of epinephrine there is a prompt rise in the cyclic AMP level of the heart which precedes the increase in phosphorylase and inotropism (Robison et aZ., 1965; Williamson, 1966) and leads to the suggestion that these last two effects are mediated by cyclic AMP. However, evidence is accumulating t o show that cyclic AMP may not play a direct role in regulating inotropism because Shanfeld et aZ. (1 969) observed an increase in inotropism without an increase in cyclic AMP. Langslet and @ye (1970) have also shown that cyclic AMP can increase phosphorylase without any increase in inotropism. These inconsistencies can be reconciled if calcium is included in the model linking the action of epinephrine with changes in heart function. As proposed by Meinertz and Scholz (1 969) and later by Rasmussen ( 1970), epinephrine may act by increasing calcium influx during the excitation process. The subsequent increase in calcium level may influence the force of contraction whereas cyclic AMP functions to stimulate metabolism. The activation process is complicated by the observation that this action of cyclic AMP on metabolism requires calcium for the conversion of phosphorylase B to phosphorylase A (Namm et aZ., 1968). Further complications may arise through the existence of feedback loops operating between cyclic AMP and calcium which closely resemble those described earlier for salivary glands (Section IV B). Calcium may inhibit adenyl cyclase (Namm et al., 1968) whereas cyclic AMP may regulate the ionized calcium level in the cytoplasm (Williamson, 1966) possibly by altering the permeability of the sarcoplasmic reticulum (Entman et al., 1969). D. EXCITATION-SECRETION COUPLING

Both cyclic AMP and calcium have been implicated in the release of a variety of cellular products such as adrenocorticotropic hormone


37

(Fleischer et al., 1969; Kraicer et al., 1969), luteinizing hormone (Samli and Geschwind, 1968; Ratner, 1970), thyroid stimulating hormone (Wilber et al., 1969; Vale et al., 1967) and amylase from the pancreas (Kulka and Sternlicht, 1968; Hokin, 1966). The way in which the various releasing factors influence the concentration of cyclic AMP and calcium and the subsequent action of these two agents in the secretory process is not clear. The control of secretion in salivary glands and the pancreas is particularly interesting since these systems are also capable of secreting large volumes of fluid. The two processes of enzyme and fluid secretion can be regulated independently of each other. For example, the submaxillary gland produces a copious watery secretion during parasympathetic (cholinergic) stimulation, while sympathetic (adrenergic) stimulation yields an enzyme-rich saliva (Bloom and Fawcett, 1968). Similarly, fluid transport by the pancreas is regulated by secretin whereas pancreozymin stimulates enzyme secretion. The way in which two or more distinct cellular activities are regulated independently of each other introduces an intriguing problem in cellular control. One possibility is that there may be a hierarchy of cellular activities which are sensitive to different concentrations of a single second messenger such as cyclic AMP. The capability of each hormone to stimulate adenyl cyclase could be different and will determine which cellular process will be stimulated. Another possibility is that each process is regulated by a different second messenger or a combination of second messengers. There is some circumstantial evidence to suggest that the latter control mechanism may operate in vertebrate salivary glands. Douglas and Poisner (1 963) showed that calcium is essential for fluid secretion during parasympathetic stimulation of parotid glands. Since there is little evidence that cholinergic agents can activate adenyl cyclase, it is conceivable that stimulation of fluid secretion is mediated by calcium but independently of cyclic AMP. However, cyclic AMP has been clearly implicated in the release of amylase by epinephrine in parotid glands (Bdolah and Schramm, 1965; Babad et al., 1967). In the case of these salivary glands, therefore, the two intracellular messengers may be independently regulated by separate primary hormones. E. CONTROL OF METABOLISM

Many behavioural patterns require instant bursts of energy and there are numerous hormonal and nervous control mechanisms which

38


assure instantaneous activation of various metabolic processes. Catecholamines are particularly important in regulating the fluctuating energy requirements of the body. Cyclic AMP can stimulate muscle and liver phosphorylase and can also activate specific lipases in adipose tissue leading to the formation of free fatty acids from triglycerides. In all the examples studied so far, however, the primary hormone can also induce changes in membrane permeability to ions in addition t o activating adenyl cyclase. Epinephrine acts on the liver t o increase the intracellular concentration of cyclic AMP which then leads to an increased release of glucose resulting from the activation of phosphorylase. The efflux of glucose precedes the increase in cyclic AMP levels (Sutherland and Robison, 1966). However, this efflux of glucose is, in turn, preceded by an efflux of potassium and calcium (Craig and Honig, 1963; Finder et al., 1964; Friedman and Park, 1968; Friedman and Rasmussen, 1970). The increased efflux of potassium and calcium probably accounts for the marked hyperpolarization of liver cells induced by catecholamines or glucagon (Haylett and Jenkinson, 1969; Friedman et al., 197 1). However, more information is required before the relationship of the ionic changes can be correlated with the hyperglycemic action of cyclic AMP. Cyclic AMP mediates the lipolytic action of various hormones by activating a specific lipase (Butcher et al., 1965; Steinberg, 1966). Apart from this metabolic effect induced by cyclic AMP, there is also evidence that lipolytic agents can induce changes in membrane potential and ion flux resembling those just described for the liver. Norepinephrine produces a marked depolarization and a rapid efflux of potassium whereas cyclic AMP and theophylline has no effect (Girardier et al., 1968; Horowitz et al., 1969; Krishna et al., 1970). Even though theophylline had no effect on the potential it was capable of stimulating heat production suggesting that the changes in membrane permeability preceded the activation of adenyl cyclase which is similar to the sequence of events observed in salivary glands (Section V C). A relationship between cyclic AMP and calcium is clearly evident in the action of parathyroid hormone (PTH) on the metabolism of kidney cells. PTH can activate renal adenyl cyclase (Chase and Aurbach, 1968) and calcium influx into kidney cells in tissue culture (Borle, 1968). As observed in salivary glands (Section IV A), the activation of adenyl cyclase by the primary hormone can occur independently of calcium (Nagata and Rasmussen, 1970). However,


39

calcium is clearly required for the ability of both PTH and dibutyryl cyclic AMP to stimulate renal gluconeogenesis suggesting that calcium is required for the expression of the cyclic AMP effect (Nagata and Rasmussen, 1970). F. TRANSPORTING EPITHELIA-THE TOAD BLADDER

Water and sodium transport by the bladder of the toad (Bufo marinus) are stimulated by the hormone vasotocin. The effects of vasotocin are mimicked by other octopeptide hormones such as vasopressin (ADH) and oxytocin. These hormones also effect the water permeability and sodium transport in other transporting epithelia such as frog skin and kidney tubules (Orloff and Handler, 1967). Calcium and cyclic AMP appear to be intimately involved in hormone action on all these tissues. Orloff and Handler (1 962) have shown that cyclic AMP and theophylline will mimic the effects of ADH on both the osmotic flow of water and sodium transport and have suggested that cyclic AMP is the intracellular mediator of the effects produced by ADH. Later they demonstrated that ADH and theophylline increase the intracellular concentration of cyclic AMP (Handler et al., 1965). Although cyclic AMP is the suggested intermediary in both actions of ADH the situation is complicated by the fact that the two effects of the hormone can be dissociated from each other yet cyclic AMP is postulated as mediating both actions. Different analogues of vasopressin have quantitatively different effects on water permeability and sodium transport (Bentley, 1967). This is suggestive of two different receptors and is seen in other amphibian epithelia (Elliot, 1967; Bourguet and Morel, 1967), In toad skin, arginine-vasotocin has a potency equal to argininevasopressin on water permeability yet is 200-400 times more potent than arginine-vasopressin on sodium transport. The effects on water and sodium movements can also be dissociated by using calcium. Increasing the serosal calcium concentration has no effect on sodium transport or stimulation by ADH, yet the effect of ADH on osmotic flow is inhibited (Petersen and Edelmen, 1964). This latter effect seems to be competitive at the receptor as it can be overcome by higher doses of ADH and as the response t o externally applied cyclic AMP is unaltered. Our understanding of toad bladder is further complicated by the presence of several cell types (Choi, 1963). The most predominant

40


cell type is the granular cell. The action of ADH on water movement appears to be at the mucosal surface of these cells as changes in the volume of these cells are seen during stimulation of osmotic water movement (Peachey and Rasmussen, 1961;Di Bona et al., 1969; Jard et al., 197 1). The situation regarding sodium transport is less clear. Sodium transport probably occurs through the mitochondria-rich cells and/or the granular cells. The latter cell type is the more plentiful but the mitochondria-rich cells appear to be more suited to active transport-they have many mitochondria and have microvilli thus resembling the parietal cells of the stomach (Choi, 1963). No changes are seen in either cell type when sodium transport is stimulated (Jard et al., 1971). Frazier suggests that the granular cells participate in ADH stimulated sodium transport whilst the mitochondria-rich cells are more suited to stimulation by aldosterone. If the granular cells are the sites of sodium transport then any theory has to explain the differences in the receptors responsible for the triggering of water and sodium movements in the same cell using the same common mediator, cyclic AMP. However, if the mitochondrialrich cells are the site of sodium transport (as suggested by Choi (1 963) and Matty and Guinness (1 964)) then separate cells would be responsible for the two effects of ADH and the explanation is somewhat easier. The receptors could have different pharmacological properties yet use similar intracellular mediators. Calcium plays an important role in the function of the toad bladder. It is important for the maintenance of the epithelial structure of the bladder and also for the action of ADH (Bentley, 1959, 1960). Removal of calcium causes the cells to dissociate from one another with a consequent reduction in transepithelial resistance and sodium transport (Hays et aL, 1965). Strontium, magnesium and barium will substitute for calcium in this respect but they will not substitute for calcium in stimulation by ADH. As well as being involved in the structural integrity of the bladder calcium may also be involved more directly in the action of the hormone. ADH will increase the efflux of 45Ca from preloaded tissue (Thorn and Schwartz, 1965) and will displace calcium from monomolecular films (Kafka and Pak, 1969). The implication of calcium involvement is also indicated from work with prostaglandins which are active in the toad bladder (Lipson and Sharp, 1971). These substances may well effect the distribution of calcium (Ramwell and Shaw, 1970). As adenyl cyclase prepared from toad bladder is inhibited by calcium (Hynie and Sharp, 1971) the same feedback mechanisms put forward


41

in this review may function too in this tissue. Therefore both calcium and cyclic AMP are involved in bladder function but their respective roles have yet to be fully ascertained.

VII. CONCLUSION

Comparison of the mode of action of 5-HT in Calliphora salivary glands with that of many other hormones reveals numerous similarities. External chemical stimuli responsible for regulating cellular activity may thus share a common mode of action. The various differences which are apparent today may represent modification of this basic mechanism developed during the course of evolution to adapt cells systems t o new environments. The main feature of this control system concerns the transduction of hormonal information arriving at the cell into internal chemical signals which can be recognized by the various effector mechanisms responsible for carrying out the actions of the hormone. In the original cyclic AMP-hypothesis the hormonal signal input was transduced into cyclic AMP through an activation of the membrane-bound enzyme adenyl cyclase and cyclic AMP alone then mediated all further cellular events. The current hypothesis proposed by Rasmussen (1970) and verified in these salivary gland studies is that the conformational changes induced in the membrane by successful hormone-receptor interactions can be transduced into more than one secondary message. There is a general labilization of the plasma membrane resulting in changes in ion permeability together with activation of adenyl cyclase. The ensuing increase in cyclic AMP concentration together with the redistribution of ions, such as calcium, are then responsible for carrying out the hormonal effect. The intracellular concentration of cyclic AMP and calcium may also be regulated by various feedback loops. The ability of cyclic AMP to regulate the internal calcium concentration may be an important component in the control of both excitable and nonexcitable cells. In some cases, for instance the slime mould, liver and Calliphora salivary glands, cyclic AMP appears t o lead to a mobilization of internal calcium whereas in heart and smooth muscle it may have the opposite effect. This very close relationship which exists between these two internal second messengers in a wide range of control mechanisms warrants careful consideration in future studies on hormone action.

42


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NOTES ADDED IN PROOF The following subjects have come to our attention since this review was submitted for publication. (1) T H E M O D E O F A C T I O N O F A D R E N O C O R T I C O T R O P H I C H O R M O N E (ACTH)

The ability of ACTH to stimulate adrenal steroid synthesis is mediated by cyclic AMP. This effect of cyclic AMP depends on calcium which appears to have a direct effect on the transfer of amino acids from aminoacyl transfer RNA to protein (Farese, 1971). Farese (1971) suggests that these observations support Rasmussen’s (1971) hypothesis that both cyclic AMP and calcium regulate intracellular events. (2) CYCLIC A M P IN I N S E C T S (a) The glycogen level of cockroach nerve cords is depleted by octopamine which acts by stimulating phosphorylase activity. This action of octopamine can be mimicked by cyclic AMP (Robertson and Steele, 1972). (b) In many vertebrate tissues examined so far, cyclic AMP appears to act by stimulating a specific protein kinase. Kuo et al. (1971) have studied the distribution and properties of protein kinases extracted from a range of tissues from different species. This study indicates that cyclic GMP may also be an important intracellular mediator of hormone action in insects.

REFERENCES Farese, R. V. (1971). Calcium as a mediator of adrenocorticotrophic hormone action in adrenal protein synthesis. Science 173,447-450. Kuo, J. F., Wyatt, G. R. and Greengard, P. (1971). Cyclic nucleotide-dependent protein kinases. IX. Partial purification and some properties of guanosine 3’,S1-monophosphate-dependent and adenosine 3 ,5 -monophosphatedependent protein kinases from various tissues and species of arthropoda. J. biol. Chem. 246, 7159-7167. Robertson, H. A. and Steele, J. E. (1972). Activation of insect nerve cord phosphorylase by octopamine and adenosine 3’,5’-monophosphate. J. Neurochem. (In Press).


Choline Metabolism in Insects R . G . BRIDGES A . R . C Unit o f Invertebrate Chemistry and Physiology. Department of Zoology. Cambridge. England I. I1.

Introduction . . . . . . . . . . . . . . Choline Metabolism in Vertebrates . . . . . . . 111. Nutritional Requirements of Insects for Choline . . A. Choline in Insect Development . . . . . . B . Substitutes for Choline in the Diet of Insects . . IV . Water-soluble Choline Metabolites . . . . . . . A. Acetylcholine . . . . . . . . . . . . B . Phosphorylcholine . . . . . . . . . . . C. Cytidinediphosphorylcholine (CDP-choline) . . D. Glycerylphosphorylcholine . . . . . . . . V. Lipid-soluble Choline Metabolites . . . . . . . A. Phosphatidylcholine . . . . . . . . . . B . Lysophosphatidylcholine . . . . . . . . C. Sphingomyelin . . . . . . . . . . . . VI . Enzymes Involved in Choline Metabolism . . . . . A. Cholineacetylase and Acetylcholinesterase . . . B . Enzymic Synthesis of Lipids Containing Choline . C. Hydrolysis of Phosphatidylcholine . . . . . D. Oxidation of Choline . . . . . . . . . . E . Synthesis of Choline . . . . . . . . . . VII . The Metabolic Role of Choline in Insects . . . . . Acknowledgements . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . .

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51 53 55 55 59 63 63 66 69 70 71 71

82 83 84 84 85

87 88 89 91 100 100

I . INTRODUCTION

The importance of choline in the metabolic processes of vertebrates has been established for many years . Nutritional studies with animals fed on low protein and high fat regimes have demonstrated that large amounts of choline are required in the diet. if normal growth is to be maintained and fatty infiltration of the liver prevented . This nutritional requirement is not. however. an 51

52

R. G. BRIDGES

absolute one and is greatly influenced by the protein content of the diet because of the interrelationship of choline, methionine and serine. Choline can be synthesized by mammals provided an adequate source of labile me thyl-groups is available. Animals possessing choline oxidase can, in turn, oxidise it to betaine which can furnish methyl-groups for the synthesis of methionine. Most symptoms of choline-deficiency can be alleviated provided adequate amounts of methionine or betaine, plus homocysteine, are available to the animal. Because of its involvement in methyl-group transfer many metabolic pathways are related to choline metabolism and this makes the results obtained from nutritional studies difficult to interpret. Choline is found in almost all living cells, predominantly or exclusively in a combined form, largely in the phospholipid fraction. Quantitatively the role of choline as a constituent of the phospholipids of the lecithin and sphingosine type overshadows its other possible functions. It is well documented that in this form it is an important component of cell membranes and that cholinecontaining lipids are the major phospholipid types in vertebrates. Qualitatively , however, choline esters play a fundamental role in neurophysiology. In vertebrates, acetylcholine plays a vital role in preganglionic transmission in the automatic nervous system, at nerve endings of the parasympathetic system, at skeletal neuromuscular junctions and in the brain. In insects, these various aspects of choline metabolism have been looked at in differing detail. The cholinergic system has received a great deal of attention because of its importance in relation to the mode of action of organophosphorus and carbamate insecticides. The composition of insect phospholipids has been analysed and results from a large number of species have accumulated over the past few years. The present review will consider only the choline-containing lipids, but in almost all cases a fairly complete analysis of the different phospholipid types has been carried out and this may be obtained by consulting the appropriate reference. A number of studies have been made following the change of phospholipid pattern during development but little has been achieved in understanding the metabolic role of the phospholipid. The need for choline in the growth of insects has been examined by dietary means in a large number of species but the replacement of this need by other related compounds has received less attention. The discovery that carnitine is an essential dietary component for tenebrionid beetles led to an examination of its ability to substitute for choline in the diet of

CHOLINE METABOLISM IN INSECTS

53

other insects. This in turn revealed that other choline-like compounds could spare the insect’s dietary need for choline and that these were incorporated into phospholipids in a similar manner to choline itself. These findings raised several important questions. How essential was the normally occurring phosphatidylcholine to the functioning of the insect? Could neurotransmitter substances other than acetylcholine be successfully used by the insect? What was the relationship between the choline involved in lipid metabolism and in acetylcholine synthesis? One of the objects of this review is to see if these questions can be answered. In doing so much of the scattered information available on choline metabolism has been gathered together. After summarizing the evidence for the insects’ need for choline and the ability of other compounds to satisfy this need, the review will deal with the occurrence of the different choline metabolites and the enzymes involved in their metabolism so that the significance of their roles may be assessed. Initially, however, an outline of the different metabolic pathways involving choline which have been demonstrated in vertebrates will be given. This will illustrate their interrela tionship and provide a basis for comparison with those found in insects. 11. CHOLINE METABOLISM IN VERTEBRATES

Figure 1 illustrates the main metabolic pathways in which choline plays a key role in vertebrates and how these pathways are Acet ylchdine

f 3 . GIycer ylphosphory ICh

y lC h

y

a

t

T

i

Betaine aldehyde

d y lCh Betaine

Sphingmyelin

PhasphatidylCh Methionine

N-Dimethylglycine

S-Adenosylmethionine

-

Phosphatidylethanolarnme

Phosphatidylserme

Fig. 1. Metabolism of choline in vertebrates.

54

R. G. BRIDGES

interlinked. Four main pathways are involved. First, the choline to acetylcholine cycle which makes up the cholinergic system involving acetyl CoA, cholineacetylase and cholinesterase (Hebb and Morris 1969). Secondly, the synthesis and catabolism of phosphatidylcholine. This involves a complex of enzymes. Choline is phosphorylated with adenosine triphosphate and cholinekinase (Wittenberg and Kornberg, 1953) and the phosphorylcholine reacts with cytidine triphosphate and choline phosphate cytidyltransferase yielding cytidinediphosphorylcholine (Kennedy and Weiss, 1956). The latter acts as the precursor for both phosphatidylcholine and sphingomy elin by transferring its phosphorylcholine moiety to a diglyceride involving 1,Zdiglyceride cholinephosphotransferase (Kennedy and Weiss, 1956) or to a ceramide involving ceramide cholinephosphotransferase (Sribney and Kennedy, 1958). Hydrolysis of phosphatidylcholine back to choline is a three step process. One of the esterified fatty acids is first removed by the action of a phospholipase A (Rimon and Shapiro, 1959) followed by the removal of the second by phospholipase B (Dawson, 1956a) and the glycerylphosphorylcholine produced finally hydrolysed to choline and cu-glycerophosphate by a glycerophosphorylcholinediesterase (Dawson, 1956b). Sphingomyelin is apparently hydrolysed back to a ceramide and phosphorylcholine (Kanfer et al., 1966). Thirdly, choline can be oxidised to betaine via betaine aldehyde involving choline dehydrogenase (Redina and Singer, 1959) and betaine aldehyde dehydrogenase (Rothschild and Barron, 1954). One of the methyl-groups of the betaine is transferred to homocysteine to give methionine by betaine homocysteine methyltransferase (Klee et aL, 1961) and the dimethylglycine further demethylated to glycine (Handler et al., 1941). Glycine and serine are inter-convertible by a serine transhydroxymethylase (Blakley , 1960) and serine can exchange with the ethanolamine-moiety of phosphatidylethanolamine to give phosphatidylserine which can be decarboxylated back to phosphatidylethanomine (Borkenhagen et al., 196 1 ). Fourthly, choline can be synthsized from ethanolamine by stepwise methylation (Bremer and Greenberg, 1961). This only occurs when it is attached to the phosphate-moiety of the phospholipid molecule. This methylation is performed by S-adenosylmethionine which is synthesized from adenosine triphosphate and methionine with the enzyme methionine adenosyltransferase (Cantoni, 1951). It should be noted that the sequence of reactions from serine to phosphatidylcholine does not lead to a net increase in phospholipid but merely an alteration in the


55

base moiety. However, ethanolamine is released on serine exchange with phosphatidylethanolamine and this is available for incorporation de novo into phospholipid after phosphorylation and formation of cytidinediphosphorylethanolamine in a series of reactions analogous t o the synthesis of phosphatidylcholine. Thus many of the compounds involved in these sequences are interrelated although some such as methionine, glycine or serine are participants in reactions other than those directly concerned with choline metab olism. 111. THE NUTRITIONAL REQUIREMENTS OF INSECTS FOR CHOLINE A. CHOLINE IN INSECT DEVELOPMENT

Insects which have been shown t o need the addition of choline to a basal diet if maximum growth of their juvenile stage is t o be achieved, are listed in Table I. In many of the early experiments casein has been used as an amino acid source. The presence of trace amounts of choline in this casein cannot be ruled out, so that the omission of choline may merely have reduced that available to the insect to some undetermined amount. Again many of the early experiments were not carried out under asceptic conditions so that micro-organisms and symbionts may have supplied some of the insects’ choline. This was well demonstrated with Lasioderma serricorne and Sitodrepa panicea which were shown to have a very definite requirement for choline in the larval diet for normal growth when freed of symbionts (Pant and Fraenkel, 1954). In spite of these provisos, the list of insects showing a requirement for choline is impressive. An attempt has been made to summarize the effects of choline deficiency on growth in Table I. The value of this may be suspect mainly because of the unknown amounts of choline present in some diets. For example, more recent experiments with Tenebrio molitor (Bieber and Monroe, 1969) have indicated that the effect of choline deficiency is much more marked in this insect than had previously been reported (Fraenkel et al., 1950). However, the effect on growth seems particularly critical in members of the Orthoptera and Lepidoptera where omission of choline from the diet results in practically no growth of the juvenile stage. Reported effects with the Coleoptera and Diptera are more variable. Many of the Coleoptera fail to grow on diets lacking choline but others merely exhibit slower

Table I Choline requirement for growth and development of juvenile stages of insects Order Coleoptera

Species Attagenus spp. Cardophilus hemipterus Dermestes vulpinus Lasioderma serricorne Palorus ratzeburgi Ptinus tectus Stegobium paniceum Tenebrio molitor Tribolium confusum

Diptera

Trogoderma granarium Aedes aegypti

Calliphora ery throcephala Cochliomyia hominivorax Drosophila melanogaster

Effect of omission of choline from diet No larval growth No larval growth Slow growth, some pupae obtained No growth in absence of symbionts No larval growth Growth retarded. Few adults obtained Growth retarded in absence of symbionts Some retardation of growth. More than 95% mortality of larvae Retardation of growth

Reference Moore (1946) Stride (1953) Fraenkel(l95 1a) Blewett and Fraenkel (1 944) Pant and Fraenkel(l954) Cooper and Fraenkel(l952) Fraenkel and Blewett (1 943) Blewett and Fraenkel (1944) {Pant and Fraenkel(l954) Fraenkel et al. (1 950) Bieber and Monroe (1 969)

Lemonde and Bernard (1 955) Fraenkel and Blewett (1943) Pant (1 956) No development Larvae reached 3rd. instar. Lea and DeLong ( 1956) Poor larval growth. Some adults. Singh and Brown (1957) Some larvae reached Akov (1962a) 3rd or 4th instar. Slow growth Sedee (1953, 1958) No larval growth Gingrich (1 964) No larval growth. No Hinton et a/. (1951) pupae. Retarded growth Reached 3rd instar. No pupae

Drosophiln subobscura

Drosophila ambigua Drosophila obscura Drosop hila funebris Drosophila immigrans Hylemya antiqua Musca domestica

Phormia regina

Pseudosarcop haga af f inis Hemiptera

Myzus persicae

Lepidoptera

Agro tis orthogonia Anthonomus grandis Argyrotaenia velutinana

Larvae reached 3rd instar. Few pupae. No affect on larval growth Mortality at pupation. Retarded larval growth. Few pupae. Near normal larval growth. Choline required for optimal growth. Poor larval growth. Some larvae reached 3rd instar. Retarded growth. Adults obtained Very retarded larval growth. No pupae

Tineola bisselliella Acheta domesticus Blattella germanica

100% mortality before 3rd instar Very retarded growth Retarded growth. Some pupae and adults Retarded growth. No pupae Very retarded larval growth. N o pupae Larval growth markedly retarded. Retarded growth Very retarded growth Very retarded growth

Locusta migratoria Schistocerca gregaria

Very retarded growth Very retarded growth

Bombyx mori Ephestia elutella Ephestia kuehniella Heliothis zea Pectinoph or gossypiella

1

Pyrausta nu bialis Orthoptera

Royes and Robertson (1 964)

Choline required for normal growth of larvae but less than for D. melanogaster Friend and Patton (1 956) House and Barlow (1 95 8) Bridges et al. (1965) Brust and Fraenkel ( I 955) McGinnis et al. (1956) Hodgson et al, ( 1 956, 1960) House (1954) Dadd e t al. ( 1 967) Kastings and McGinnis (1 967) Vanderzant (1 963) Rock (1969) Horie and Ito (1 965) Fraenkel and Blewett (1946b) Vanderzant (1968) Vanderzant and Reiser (1956) Beck et al. (1949) Fraenkel and Blewett (1 946a) Ritchot and McFarlane (1961) Noland and Baumann (1949) Noland et al. (1949) Dadd (1961) Dadd (1961)

58

R. G. BRIDGES

growth. With the Diptera the reported effects range from no larval growth for Cochliomyia hominivorax to near normal growth of larvae for Musca domestica and Phoimia regina. The report by Singh and Brown (1 957) that adults of Aedes aegypti were obtained, was due almost certainly to the presence of lecithin which was included in the diet. It seems likely that some Diptera require relatively small amounts of choline when compared with other orders and that the variations in the observations are due to trace contamination of choline in the other compounds of the diet. For example, the addition of as little as 0.25 pmole of choline/g of diet is sufficient to enable housefly larvae t o develop to give adults (Bridges and Ricketts, 1968) whereas at least 5 pmole/g of diet is required for the normal development of Bombyx mori (Horie and Ito, 1965). The need for choline by Hemiptera is not clearly established. Choline deprived diets affected the growth of Myzus persicae but adults were obtained, although in reduced numbers. The black citrus aphid, Toxoptera aurantii, could be reared for at least two generations on an amino acid diet lacking all B vitamins (Tahori and Hazan, 1970). However, the authors suggested that this was due to the presence of the necessary symbionts in the aphid. The need for a dietary supply of choline by the adult insect has been studied in only a few cases. Some insects do not feed during their adult life and others such as the housefly will live on a diet of glucose and water but will not lay eggs. Rasso and Fraenkel (1 954) showed that the addition of choline to the diet of adult female blowflies, Phormia regina, somewhat accelerated egg production. Similarly Vanderzant and Richardson ( 1964) concluded that the adult Anthonomus grandis required choline in the diet for continued laying of viable eggs, although choline had been supplied in the larval diet. The utilization of dietary supplied choline by the adult Drosophila melanogaster has been quantified by Geer et al. (1 970). Choline is required for both egg and sperm production by this insect. If it is not supplied to the adult female some eggs will be laid at the expense of the insect’s reserves of choline. Thus at least in the few orders which have received most attention, choline is required by the insect for normal growth. Differences recorded in the effects of deficiency may be due t o variation in the needs of the insect combined with the presence of trace amounts of choline contaminants in the diet. From the limited amount of information available, it seems likely that once the adult stage is reached, the insects main need for choline is for reproduction.


59

B. SUBSTITUTES FOR CHOLINE IN THE DIET OF INSECTS

Table I1 summarizes information available on the ability of various compounds related to choline to act as a substitute for an established need for choline in the diet of different insects. All diets contained methionine either as the free amino acid or as a constituent of the casein used as an amino acid source and any relationship between the choline and methionine content of the diet has not been studied in detail. Apart from work with Blattella germanica, betaine has not been found to be a substitute for choline in the diets of the insects examined. In the case of the cockroach, the sparing action of betaine may have been due to the presence of internal symbionts. Betaine aldehyde has been shown to be an effective dietary substituent for choline when fed at high concentrations to Drosophila melanogaster. Addition of ethanolamine has failed to stimulate the growth of any of the insects tested. N-Monomethylaminoethanol has some slight stimulatory effect on growth with the exception of Musca domestica where larval growth was found to be more retarded. N-Dimethylaminoethanol stimulated growth of larvae and pupation occurred in Phormia regina, Drosophila melanogaster and Musca domestica. No adults were obtained however. The same compound also stimulated growth of Tribolium confusum, Aedes aegypti and Blattella germanica, although in the latter case it was fed along with methionine. This stimulation by dimethylaminoethanol has led some authors to suggest that synthesis of choline from this precursor may be occurring. However, it has been shown (see Section VA) that this compound can be incorporated into the phospholipids of Phormia regina and Musca domestica and this may well be connected with its growth stimulating property in these and other insects. Considerable attention has been paid to carnitine as a possible substituent for choline in insect diets, although there is no evidence that it will act as a choline substitute in mammals (Fitz and DuPont, 1957). Carnitine has been established as a dietary essential for the normal development of Tribolium confusum (French and Fraenkel, 1954), Tribolium castaneum (Magis, 1954), Tenebrio molitor and Palorus ratzeburgi (Fraenkel, 195 1b; Cooper and Fraenkel, 1952). However, there is no evidence that it would replace the choline requirements of Palorus ratzeburgi, Tenebrio molitor, Blattella germanica, Bombyx mori, Anthonomus grandis, Aedes aegypti or Cochliomylia hominiuorax. It would, however, act as a partial replacement for choline in the diets of larvae of three Diptera,

Table I1 The effectiveness of various compounds as substitutes for choline in the diet of insects. Betaine

N-Dimethylaminoethanol

Lasioderma serricorne

-

-

Palorus ratzeburgi

-

N-Monomethylaminoethanol -

Ethanolamine -

Carnitine

Referencc

No substitute

Fraenkel et al. (1955)

Requires carnitine Fraenkel et al. as vitamin but will (1955) not substitute for choline Requires carnitine Bieber and as vitamin but will Monroe (1 969) not substitute for choline

Tenebrio molitor

Tribolium No substitute confusum Aedes aegypti

No substitute

Cochliomyia

N o substitute

hornin ovorax

Slight stimulation of growth Slightly enhanced growth, not stimulated with additional methionine

Very slight stimulation of growth

-

No substitute

-

N o substitute

Requires carnitine Lemonde and Bernard (1 955) as vitamin No substitute

Akov (1 962b)

N o substitute

Gingrich (1 964)

Drosophila

melanogaster

No substitute.

Increase in

S o m e increase

Betaine aldehyde effective at high concentrationsa

larval growth. No adults."

in larval growth. Not so effective as dimethylaminoethanola

Musca No substitute dom estica

Increases larval growth. Pupae obtained No adults

Retards larval growth

Anthonomus No substitute grandis Bombyx mori

-

Blattella Adequate germanica substituteC

Phormia regina

No substitute

No substitutea Effective

substitute. in smaller

-

Increase in Some increase in larval growth larval growth. Pupae obtained. No adults

(1965) Geer e t al. (1968) bFraenkel et al. (1955)

No substitute

Effective Bridges e t al. substitute. Adults (1965) obtained.

No substitute

No substitute

Vanderzant (1 963)

N o substitute

Horie and Ito (1965)

-

N o substitute but with methionine some enhancement of growthC

"Geer and Vovis

No substituteC N o substituted

No substitute

CNoland and Baumann (1 949) Fraenkel e t al. (1955)

Effective Hodgson et al. substitute. Adults (1956, 1960, obtained but not 1969) considered normal

62

R. G. BRIDGES

Phormia regina, Musca domestica and Drosophila melanogaster. With these insects larval growth was normal when carnitine was supplied in the diet in place of choline and pupation occurred. Adult flies were obtained, although in the case of Phormia regina none were considered t o be normal. The finding that carnitine was decarboxylated to 0-methylcholine which was incorporated into the phospholipids of Phormia regina larvae (Bieber et al., 1963) led to an intensive investigation into the ability of various compounds related to choline to act as dietary substitutes in the three Diptera. The results are summarized in Table 111. In general a common pattern emerges for all three insects. With Phormia regina, adults were obtained when /3 -meth ylcholine, a-methylcholine , dimethylethylcholine and dimethyl-isopropylcholine were included in the larval diet in place of choline. 0-Methylcholine and dimethylethylcholine were the most successful choline substitutes for the housefly but a-methylcholine failed t o support growth t o adult stage. 0-Methylcholine and dimethylethylcholine supported growth of Drosophila melanogaster to the adult stage. Most other choline analogues increased larval growth and enabled the larvae to pupate but very few adults were obtained. It would seem that the 1-hydroxy-2-quaternary ammonium grouping was essential for the compound to be an adequate substitute for choline. A single methyl-group substituent on the 1- or 2-position was acceptable but further substituents in the 2-position made the compounds less effective. Alkyl-substituents of the quaternary nitrogen groups were permissible if one methyl was replaced by an ethyl-group or an isopropyl-group (Phormia regina) but compounds with alkyl-groups of increasing chain length were progressively less effective although the diethyl-compound stimulated larval growth in the case of Phormia regina and yielded a few adults in the case of Drosophila melanogaster. Thus only very close analogues of choline are capable of acting as dietary substituents which enable the insect to develop to the adult stage. It is of interest to note that adults of Drosophila melanogaster (Geer et al., 1968) from larvae fed on a medium containing carnitine in place of choline and themselves fed on a similar medium laid only few eggs, none of which hatched; so that although certain choline analogues enable adult development, it is likely that choline will be required if they are to produce viable eggs. So far studies with choline analogues have been carried out almost entirely with the three related species of Diptera. The failure of carnitine to act as a replacement for choline on other insects does not imply that they are incapable of utilizing choline analogues, rather that they are unable


63

to decarboxylate carnitine t o 0-methylcholine. Bieber and Monroe (1969) have shown in fact that Tenbrio molitor larvae were unable to decarboxylate carnitine but that 0-methylcholine could act as a substitute for their choline requirement, enabling near normal growth. From the results obtained from all these nutritional studies it seems that choline, or some close analogue, is a dietary necessity for the development of insects. This would suggest that, if insects can synthesise choline, they are unable to synthesise adequate amounts to meet their requirements. Apart from the results with Blattella germanica, the failure of betaine, ethanolamine or N-methylated ethanolamines to substitute for choline suggests that transfer of me thyl-groups is lacking which implies a complete inability to synthesise choline. However, the interpretation of results from these nutritional experiments is difficult and only from a detailed investigation of the biochemistry of each insect can a definite conclusion be drawn. IV. WATER-SOLUBLE CHOLINE METABOLITES A. ACETYLCHOLINE

0

+ II (CH3)3-N-CH2 CH2 0-C-CH3 The presence of acetylcholine in insects is now well established and has been the subject of several recent reviews (Colhoun, 1963; Treherne, 1966; Boistel, 1968; Smallman and Mansingh, 1969). Acetylcholine is present in high concentrations in insects when compared with most other invertebrates and vertebrates. It is now generally accepted that it is an excitory synaptic transmitter in insect nerve (Kerkut et al., 1969) but not at the neuromuscular junction. Colhoun (1963) and Smallman and Mansingh (1969) have summarized data showing the distribution of acetylcholine in various insect tissues and at different stages of development. In Periplaneta americana, acetylcholine is confined to nervous tissue and recent unpublished work by the author has also indicated that it is largely associated with the fused ganglia of the larva of the housefly. High concentrations of acetylcholine have been reported in the eggs of insects. The most complete study has been that of Mehrotra (1960) on Musca domestica and Oncopeltus fasciatus. A large increase in the acetylcholine content of the eggs occurred just prior

Table 111 Utilization of choline analogues in place of choline by Drosophila melanogaster, Musca domestica a n d Phormia regina

D. melanogaster

M. domestica

P. regina

A (CH3)3fi-R

R=

OH

1

-CH, CHCH,

p-methylcholine

Normal L growth, P and some A 0btainedC.f

Normal L growth, P and some A ob t h e & * k

Normal L growth, P but no A obtainede

Normal L growth, P and some A obtainedg

Some enhancement of L growth. Few P and no A obtaineda

Normal L growth, some P but no A obtainede

Some enhancement of L growth. No P obtainedg

-

Normal L growth, P but no A obtainede

Enhances larval growth. P and some A obtaineda

CH, OH 1 1 -CH-CH, (YH3)2yH -C

-

CH,

a,&-dimethylcholine

C,H, OH

I

1

CH,

a-e thylcholine

-CH, CH, CH, OH

Homocholine

-C

-

Enhancement of L growth. P and some A obtaineda

R

I B (CH,),-N+-CH,CH20H

R= N-DimethylaminoEthan01

Enhancement of L growth. Some

P and few

Normal L growth, P but

Enhancement of L growth,

no A obtainedc.d

P but no A obtained8.i

C*H, -

Dimethylethylcholine

C,H, (CH,), -CHC,H, -

c5 HI,

-

Enhancement of L growth.

Normal L growth,P and

P and A obtaineda

A obtainede

Dimethyl-n-propylcholine Dimethyl-isopropyl- . choline Dimethyl-n-bu tylcholine Dimethyl-n-amylcholine

-

-

Enhanced L growth, P but very few A obtainede -

Enhanced L growth, P but very few A obtainede Some enhancement of L growth, few P and no A obtainede

Normal L growth. P and

some A obtained89h.i Enhancement of L growth, some P, no A obtainedg.ki Normal L growth, P and A obtainedg9h.i Enhanced L growth, some P but no A obtainedg.h,i -

C Miscellaneous N-Monomethylaminoethanol CH,N-CH,CH,OH 7-Butyrobetaine (CH,), - N'- CH, CH, CH, COOH Diethy lmethylcholine (CH&

Enhancement of L growth. Some P no A obtained4 b Enhancement of L growth. P and some A obtaineda

Decrease in L growth. No P 0btainedc.d -

-

Enhancement of L growth. No P obtained8 Enhancement of L growth. Some P and A obtainedgsk Enhancement of L growth. Some P but no A

I

(C,H5)l -N'CH,CH,OH Sulphocholine (CH,), - S+ CH, CH,OH

Enhancement of L growth. Some P and A obtained0

Triethylcholine (C,Hs)3- N+-CH, CH, OH

Enhancement of L growth. Few P and no A obtaineda

and Vovis (1965) b Geer et al. (1968) C Bridges et al. (1965) Bridges and Ricketts (1967) Bridges and Ricketts (1970) fMoulton et al. (1970)

a Geer

L=Larval

g Hodgson et al. (1969) h Mehendale et al. (1970) i

Hodgson and Dauterman (1964)

i Bieber and Newburgh (1963) k Bieber et al. (1963

P = Pupae

A = Adults

ob tainedg. i Enhancement of L growth. Some P but no A ob tainedg -

66

R. G. BRIDGES

to hatching, rising to 3 pmole/g in eggs of Musca domestica and 1 pmole/g in eggs of Oncopeltus fasciatus. This is very much higher than values obtained for 6-day-old larvae of Musca domestica which were between 0.01 8-0.024 pmole/g (Bridges and Ricketts, 1968, 1970). It is difficult to explain these high levels in terms of function and the source of the large amount of choline necessary for its synthesis is not at all clear. From the data collected by Smallman and Mansingh (1969) little change in the acetylcholine content is observed during the late larval and pupal stages of Antheraea polyphemus, Antheraea pernyi, Malacosoma americanum, Galleria mellonella, Ostrinia nubilalis and Hyalophora cecropia. During development of the adult, a rise occurs which appears to be associated with the differentiation and elaboration of the imaginal nervous system. Colhoun (1963) has noted that there are some events, remote from the insect nervous system, that involve acetylcholine. For example a high concentration has been reported in royal jelly of Apis mellifera and also in the venom of Vespa crabro. It is also present in the male and female reproductive organs of certain Lepidoptera. A recent paper by Grzelak et al. (1 970) suggests that changes observed in the acetylcholine content of the abdomen of Celerio euphorbiae during the first eight days after pupation do not seem to involve the central nervous system. Thus although insects have a basic need for choline to provide acetylcholine for the normal functioning of the nervous system some may have an additional need to supply the acetylcholine required for more specialized purposes. B. PHOSPHORYLCHOLINE

+

0 II

(CH3)3N CHZCHZ-0-P-OH

I OH Phosphorylcholine has been found in high concentrations in the haemolymph of certain Lepidoptera. Table IV summarizes the information available giving the concentration of phosphorylcholine in the lymph or tissues and its percentage of the acid-soluble phosphorus. The presence of undetermined amounts of phosphorylcholine has been demonstrated in adult Celerio euphorbiue (Chojnacki, 196 l ) , adult Arctia caia (Chojnacki and Korzybski,

67


Table IV Phosphorylcholine content in insects

pmole/mlOr g wet wt. Musca Whole L (6 day-old) dornesticaalb Whole A Antheraea Haemolymph P, 5 months at 6zC polyphemusC P, 9 months at 6 C P, 11 months at 6°C Bombyx morF Haemolymph L, midstar V L, instar V pooled P, early Galleria Haemolymph L, Days fasting 0 mellonellad Days fasting 5 Days fasting 19 Days fasting 37 Days fasting 42 Galleria Intestine L mellonellae Fat body L Remaining tissue L Whole L Hyalophora Haemolymph L, instar V cecropiaC P, 25OC A? 2 days old A? 13 days old Hyalophora Fat body P, diapausing cecropiaf P, 2 days after start of adult development Protoparce Haemolymph L, fully grown sex tac qb Bridges and Ricketts (1968,1970)

dLenartowicz and Niermierko (1964) fGarey and Wyatt (1963) L = Larva

P = Pupa

% of acid soluble phosphorus

0.19-0.45 0.84 10.7 11.1 15.6 6.7 5.7 18.0 21.2 24.6 35.4 28.6 43.9 6.2 4.1 5.9 11.7 2.3 11.5 8.7 20.7 0.4

48 33 47 13 15 62 29 36 48 35 57 14 20 17 35 8 31 24 64 18

1.7 w.7

25 3-3

Wyatt e f al. (1963) Lenartowicz et a1. (1964) A = Adult

1962), larvae of Heliothis zea (Willis and Hodgson, 1970), in haemolymph of Acan tholyda nemorulis larvae (Zielinska and Dominas, 1967), larvae of Hylobis pales (Willis and Hodgson, 1970), adult Periplaneta umericana (Kumar, 1968) and in larvae of Phormia regina (Willis and Hodgson, 1970; Mehendale et al., 1970). Phosphorylated choline-analogues have been demonstrated in Phorrnia regina larvae when fed on diets containing dimethylethyl-

68

R. G. BRIDGES

choline, dimethyl-n-propylcholine, dimethyl-isopropylcholine and dimethyl-n-butylcholine (Mehendale et al., 1970). Phosphorylcholine was found in extracts of Musca dornestica larvae and phosphoryl-0methylcholine when 0-methylcholine, carnitine or y-butyrobetaine were fed (Moulton et al., 1970; Bieber et al., 1969). The concentration of phosphoryl-p-methylcholine in these larvae was estimated to be between 0.3 and 0.5 pmole/g wet weight. Estimates of the amount of phosphorylcholine have been made in larva and adults of Musca domestica (Bridges and Ricketts, 1968, 1970) and it has been shown to depend on the amount of choline available to the larvae in its diet. The figures given in Table IV are for housefly larvae fed on high choline diets (16 pmole choline/g casein). When fed on diets containing 1 pmole and 0.04 pmole choline/g casein the levels of phosphorylcholine in the larvae fell to 0.02 and 0.001 pmole/g wet weight respectively. Phosphorylcholine was not reported as a component in the phosphorus-containing compounds present in the acid-soluble fraction of thoraces of Musca dornestica (Winteringham et al., 1955). However, using their figure of 10 pg acid-soluble phosphorus/7 mg thoracic tissue, the amount of phosphorylcholine present in the adult fly (Table IV) would be only 2% of the total phosphorus-containing compounds and could well have remained undetected. The low proportion of phosphorylcholine in this insect is very different from the figures obtained for the Lepidoptera (apart from Protoparce sexta), where even in tissue extracts as distinct from haemolymph, about 20% of the acid-soluble phosphorus was phosphorylcholine. The high levels of phosphorylcholine in the haemolymph of the Lepidoptera listed in Table IV were found at all stages of development there being some suggestion of an increase from larva to adult. It was particularly high in larvae of Galleria mellonella and it increased during fasting. Lenartowicz and Niemierko (1964) suggested that this was due to the breakdown of phosphatidylcholine. This implies the action of a phospholipase C or some factor affecting the reincorporation of phosphorylcholine into phospholipid such as a lower availability of &glyceride. No role has been suggested for the high concentrations of phosphorylcholine in the haemolymph of these insects. Whether they are related to the amount of choline in the diet is not known with certainty but the high level found in fasting Galleria mellonella larvae suggests that they may be independent. Whatever the role of this compound is in these insects, choline from the diet will be required for its synthesis. In summary, the presence of phosphorylcholine has been

69


demonstrated in a number of different orders of insects; the Coleoptera, Diptera, Hymenoptera, Lepidoptera and Orthoptera. The amounts present show some dependence on the stage of development and, in the case of the housefly, on the amount of choline available to the insect in its diet. Tissue levels in most cases are comparable with those reported for vertebrates (Ansell and Hawthorne, 1964) but the concentration in the haemolymph of the Lepidoptera studied (apart from Protoparce sexta) is very much higher. C. CYTIDINEDIPHOSPHORYLCHOLINE (CDP-CHOLINE)

R B

CH2-0-P-0-P-0-CH2CH2 I

OH

4.

N(CH3)3

I

OH

Very little information is available on the occurrence of this choline metabolite in insects. It is the intermediate between phosphorylcholine and the lipids phosphatidylcholine and sphingomyelin. In vertebrates the reported levels of CDP-choline do not exceed 0.1 pmole/g (Ansell and Hawthorne, 1964). Results from feeding isotopically labelled choline have shown the presence of CDP-choline in larvae of Phormia regina (Mehendale et al., 1970) and in larvae and adults of Musca domestica (Bridges and Ricketts, 1968, 1970). In the case of the housefly, the level of CDP-choline depended on the level of choline in the larval diet and was 0.006 pmole/g wet weight of tissue (dietary concentration of choline 16 pmole/g casein) 0.002 pmole/g ( 1 pmole/g casein) and 0.0003 pmole/g (0.04 pmole/g casein). Adult flies from larvae fed on diets containing 16 pmole choline/g casein contained 0.009 pmole CDP-choline/g of fly. The formation of CDP-choline has been demonstrated in vitro in fat body preparations from larvae of Phormia regina (Shelley and Hodgson, 1970). Willis and Hodgson (1970) stated that CDP-choline could be detected in extracts from larvae of Heliothis zea and Hylobius pales as well as Phormia regina. Thus, where it has been looked for in insects, CDP-choline has been found. In the housefly larva, the concentration of CDP-choline is dependent on the amount of choline in the diet and appears to be considerably lower than that found in vertebrates.

70

R. G. BRIDGES

D. GLYCERYLPHOSPHORYLCHOLINE

CH2OH

I

CH OH

I

OH Glycerylphosphorylcholine can be produced by the action of phospholipases on phosphatidylcholine. It has been found in fairly high concentrations in the fat body of the pupae of Hyalophora cecropia (Garey and Wyatt, 1963). Diapausing pupae, one month after pupation stored at 25-30" contained 0.1 1 pmole/g (5% of the acid-soluble phosphorus); diapausing pupae, debrained two months after pupation and stored at 25" for three months and then at 6" for three months contained 4.65 pmole/g (40% of the acid-soluble phosphorus) and pupae at second day of adult development after storage for seven months at 6°C and two days at 25°C contained 0.68 pmole/g (1 0% of the acid-soluble phosphorous). Wyatt et al. (1 963) failed to detect glycerylphosphorylcholine in the haemolymph of Hyalophora cecropia but they suggested that this might be due to its hydrolysis to a-glycerophosphate and choline during storage of the lymph prior to analysis. Kumar ( 1968) found injected methyl-14C-choline appearing in glycerylphosphorylcholine in Periplaneta americana. Mehendale et al. ( 1970) found radioactive glycerylphosphorylcholine in extracts from Phormiu regina larvae fed on diets containing methy1-l4C-choline. Unpublished results by Bridges have demonstrated very small concentrations of g1ycerylphosphorylcholine in housefly larvae of the order of 0.01 pmole/g wet weight. Glycerylphosphoryl0-methylcholine has been detected in housefly larvae fed on diets containing 0-methylcholine, carnitine or y-butyrobetaine (Bieber et al., 1969). The presence of large amounts of glycerylphosphorylcholine in fat body of Hyalophora cecropia during pupation may be due to breakdown of phospholipid during this stage of development. However, the amount which accumulates is too great to be entirely derived from pre-existing phospholipid and suggests a steady


71

turnover of phospholipid leading to this product. The presence of this choline metabolite has been detected also in Orthoptera and Diptera but apparently in much smaller amounts. V. LIPID-SOLUBLE CHOLINE-METABOLITES A. PHOSPHATIDYLCHOLINE

A

CH, -0-C-R

I

‘ 1

CH-0-

-R”

OH R‘ and R” = long chain alkyl groups Table V lists the percentage of the phospholipid present as phosphatidylcholine in lipid extracts from different orders of insects at different stages of development. It is an extension of the major survey carried out by Fast (1966) and adds further confirmation to his conclusion that, as in vertebrates, phosphatidylcholine is the predominant phospholipid type in Hymenoptera, Lepidoptera and Orthoptera. The Coleoptera, Hemiptera and the one species of Thysanura examined contain a rather lower percentage of their phospholipids as phosphatidylcholine. The Diptera, apart from a few exceptions (Ceratis capita eggs and larvae, Culex pipiens fatigans eggs, Glossina morsitans adults, Phytophaga rigidae larvae and Rhabdophaga swainei larvae) all contain a much lower level of phosphatidylcholine than other insects and vertebrates, varying between 14 to 29% of the total phospholipid. The figure of 22% reported for larvae of Trogoderma granarium is the only other exception to these generalizations. Tables VI and VII summarize the

4 N

Table V Lipids containing choline in whole insects Results are based on phosphorus estimation and are expressed as a percentage of the total lipid phosphorus

Coleoptera Altica am biens alni Chrysomela crotchi Pissodes strobi Tenebrio molitor Tribolium confusum Trogoderma granarium Diptera Aedes aegypti Calliphora ery throcephala Ceratis capita

Stage

Phosphatidylcholine

L L P L L L P A L A

34.3 39.8 36.0 39.3 43 42-53 39-44 35-36 22.2 37.7

L L E

14.4 24 31.8 34.3 26.0 17.5 26.1 33 26 23 20 27.5 27.4 14.9 Present

? ? 0.1 1.9 1.o 1.1 1.8 ? ? ? ? 1.8 0.4 1.5

17.5

?

L Chironomus spp. Culex pipiens fatigans

Dacus oleae

P A A

E L P A L P A

Drosophila melanogaster iigy&1s.itB~ikTsa

A L

Lysophospha- Sphingomyelin tidylcholine ? ? ? ? ?

trace 1-7 7

? ? ? ?

Reference

Fast (1 966) Fast ( 1966) Fast (1966) Fast (1966) Kamienski et al. (1 965)

6.8

9 Beaudoin et al. (1 968) 9-10

1.3a Absent Absent >

Fast and Brown (1 962) Bridges (unpublished work)

} Castillbn et al. (1971)

Absent Absent Absent Absent Absent >

Absent Absent Absent) Absent Absent Absent ? Absent

W

E

U

Rao and Argarwal(1969)

?

P

}

I

Fast (1 966) Kalra et al. (1969)

Castillbn et 41. (1 971) Wren and Mitchell (1959) Fast (1966)

8

v1

Glossina morsitans Hylemia antiqua Lucilia cuprina Musca domestica Nephrotoma sodalis Phormia regina Phytophaga rigidae Rhabdophaga swainei Strauzia longipennis Hemipt era Anuraphis bakeri Aphrophora parallela Ericerus pela Oncopeltus fasciatus Prociphilus tesselatus Schizolachnus pini-radiatae Hymenoptera A can tholyda nemoralis Arge pectoralis Monoctenus juniperinus Neodiprion sertifer Pikonema alaskensis Lepidoptera Archips cerasivoranus Arctia caia Bom byx mori Datana integerrima Erannis tiliaria

A

A P&A L A A L E L A L L L

Au N.A L E,N,A A All All

17.4 26.0 16.0 13.9 17.2 26.0 19 21 19 37.5 48.1 29.0

Absent ? ? ? ? ? ? ? ? ?

27.2 46.0 30 Major phospholipid 31 29.7 33.7

L L L L L

64.5 41.6 52.1 49.2 47.8

L A

45.9 47

P L L

62.5 37.6 45.4

Absent Absent Absent Absent Absent Absent Absent Absent Absent Absent Absent Absent Absent

? ? ?

35

-

? ?

? ? ?

Cmelik et aL (1 969)

Fast (1 966) D'Costa and Birt (1966) Bridges and Price (1 970a) Fast (1 966) Crone and Bridges (1963) Fast (1966) Bieber et al. (1 96 1) Fast (1 966) Fast (1 966) Fast (1 966)

?

?

7

7

Fast (1 966) Fast (1966) Hashimoto and Mukai (1 967) Kinsella (1 966a) Yurkiewicz and Whelchel(l969) Fast (1 966) Fast (1 966)

? ? ? ?

? ? ? ?

Zielinska and Domina (1 967) Fast (1 966) Fast ( 1966) Fast (1 966) Fast (1966)

? ?

1W

trace Present

Present

1

u 7.8

? ? ?

?

16" ? ?

Fast (1 966) Chojnacki and Korzybski (1 962) Sridhara and Bhat (1 965) Fast (1 966) Fast (1966)

w

4 P

-

Table V-cont.

Galleria mellonella Heliothis zea Hyphantria cunea Paleacrita vernata Plodia in terpu n c tella Protoparce sexta Orthoptera Acheta domesticus Blattella germanica Diapheromera femorata Gryllus bimaculatus Locusta migratoria Periplaneta americana

L

?

60

A 39 L & P Major component

Not detected Wlodawer and Wihiewska (1 965) 5 Yurkiewicz (1 968) Present Lambremont and Graves (1 969) ? Fast (1 966) 7 Fast (1966) Yurkiewicz (1 967) 8 Not detected Hodgson ( 1 965)

?

L L E

40 48.7 38-45

? ?

u

7 p

A

ca 40

?

W

E L

58 40 45 53 57.4 54 58.6-70.5 5 7-6 5 57.7 60 44

? ? ? ? ? ? ? ? Some detected ? ?

A A A A E

E N A A

11

}

? ?

Present 3.5-4.W 5.2-6.5 6.1 5.9 ?

}

Lipsitz and McFarlane (1 970) Crone and Bridges (1 963) Fast (1 966) Fast (1 967) Allais et al. (1 964) Kinsella (1 966b) Crone and Bridges (1 963)

Thysanura Lepisma saccharina a Sphingomyelinnot fully characterized.

4.0

36.3 E = Egg;

L = Larva;

10.4 N = Nymph;

Kinsella (1 969) P = Pupa;

A = Adult.

E U Q g

Table VI Lipids containing choline in different insect tissues Results are based on phosphorus estimations and are expressed as a percentage of the total lipid phosphorus Phosphatidylcholine

Lysophosphatidylcholine

Sphingomyelin Absent Absent Absent Absent

Reference 0

Musca domestica L Brain + imaginal discs Cuticle Muscle + tracheae Gut + salivary glands + Malpighian tubules Fat body Haemolymph A Head Thorax Abdomen Sarcophaga bullata L Fat body Apis mellifera A Brain Galleria mellonella L Haemolymph

17.7 20.6 19.1 19.5

? ? ? ?

19.2 11.3 19.5 13.5 19.0 21 12 57

? ? ? ? ? 13 ? ?

Hyalophora cecropia PA Muscle A Muscle Acheta domesticus L Haemolymph Schistocerca gregaria A Haemolymph

46 43

X 0

'I

c Bridges and Price ( 1970a)

d z m 4

>40

Absent Absent Absent Absent Absent Absent 9 Not detected

I

Crone ( 1 964)

h-

All detected but not quantified Detected Detected PA = Pharate adult;

}

sE z

2 Allen and Newburgh (1 965) Patterson et al. (1945) Wlodawer and Wiiniewska ( 1 96s)

? ?

L = Larva;

k

. - - - - I

Thomas and Gilbert (1 967)

.

Wang and Patton (1 969) Mehrotra et al. (1 966)

A = Adult.

a

c4l

m

0

2

Table VII Lipids containing choline in various subcellular fractions from insect tissue

4 o\

Results are based on phosphorus estimations and are expressed as a percentage of the total lipid phosphorus Phosphatidylcholine Lucilia cuprina Flight muscle sarcosomes PA 20 A at emergence 27 A at 5 day 21 Musca domestica Gut 600g Fraction L, 6 day 7.1 10,OOOg Fraction 14.4 100,OOOg Fraction 20.5 Muscle 600g Fraction L, 6 day 13.8 10,OOOg Fraction 11.6 100,OOOg Fraction P > 0.1) and the slow fraction showing no significant correlation (P 0.8).

296

J. E. TREHERNE AND Y. PICHON

0

5

10

time

15

(min)

Fig. 20. Semi-logarithmic plot of "Na efflux from an intact, isolated cockroach connective illustrating analysis of the efflux curve into three components: slow component (linear portion of open circle curve), intermediate component (straight portion of squares curve) and fast component (triangles). From Tucker and Pichon, 1972b.)

The highly significant correlation between the fast efflux and the size of the extraneuronal potentials agrees with the quantitative analyses of the ionic basis of this potential (p. 287). In particular, these experiments support the postulated role of the intercellular perineurial channels in short-circuiting the trans-perineurial potentials observed in P. americuna (Pichon et al., 1971) and possibly also in M. sextu (Pichon et al., 1972). In desheathed cockroach connectives 22Na efflux of all three exchange components was increased as compared with intact preparations (Tucker and Pichon, 1972b). As will be seen from Table I1 the effects were most marked in the case of the rapidly-exchanging 22Na fraction. As has been emphasized (p. 268) the perineurial damage associated with the desheathing procedure renders extensive volumes of the glial cytoplasm accessible to peroxidase molecules and also causes a dramatic increase in the measured inulin space (Treherne, 1962a). These results have also been correlated with the

297

INSECT BLOOD-BRAIN BARRIER FAST

COMPONENT

I 1

0

I

I 2 time

3

(min)

Fig. 21. Relationship between the rate of sodium efflux in the fast component (Fig. 20) and the magnitude of the potassiuminduced extraneuronal potentials in intact cockroach connectives. The line drawn through the points is the calculated regression line. (From Tucker and Pichon, 1972b)

extremely rapid ion movements measured in electrophysiological experiments in desheathed preparations which have been postulated to result from an increased accessibility of glial cytoplasm to inorganic ions contained in the bathing medium (p. 268). It, therefore, seems reasonable to assume that the substantial decrease in the measured half-time of the rapidly-exchanging 22Na fraction results, in part at least, from the rapid escape of an appreciable intracellular ion fraction. The alkylammonium cations which penetrate into the tissues of the intact abdominal nerve cord of P. arnericana can be resolved, in Table I1 The effect of desheathing on 22Na efflux from nervous connectives in Periplaneta americana * Preparation

Fast component 10.5 (sec.)

Intermediate component ro.5 (sec.)

Slow component to.5 (min.)

Sheathed

29.7-110.1

102.2k17.7

16.0f1.4

P Desheathed

2.0k0.82

* From Tucker and Pichon (1972b)

< 0.001

33.0f2.72

P

< 0.05 P < 0.02 8.5f0.47

298


radiotracer experiments, into fast and slowly-exchanging fractions (Eldefrawi and O’Brien, 1967). It was suggested that these represent the distribution of cations between extracellular and intracellular phases or conceivably, between “free” and “bound” extracellular cations. The data presented by Eldefrawi and O’Brien show that increasing lipid solubility tended to increase the penetration into the central nervous tissues and that increasing ionic size decreased the rate of influx. Charged molecules appear to penetrate more slowly than uncharged ones of equivalent dimensions, the difference being, however, relatively small (of the order of a 5 to 15-fold change). It is very difficult, in our present state of knowledge, to relate the degree of accessibility of ions of differing physico-chemical properties to the extent of their penetration within the different structural compartments of the central nervous system and, in particular, to their accessibility to the neuronal elements. It is hoped that future research may elucidate the role of factors such as lipid solubility in the penetration of substances to the various neuronal surfaces of the insect central nervous system. Our knowledge of the uptake of nutrient substances is limited to a single early investigation on the uptake of radioactive glucose and trehalose molecules by the cockroach abdominal nerve cord (Treherne, 1960, 1961a). As in the case of cations a relatively rapid uptake of 14C-labelledglucose and trehalose was demonstrated, being equivalent to an influx of 1.09 mM glucose/l nerve cord water. The possible significance of these results can be best appreciated in relation to the contemporary histochemical study of Wigglesworth (1 960) on the uptake of carbohydrate by the central nervous system of P.americana (p. 305). In spite of the apparent insensitivity of insect ganglia to externally applied acetylcholine (p. 258) relatively rapid exchanges of l4C-labe1led acetylcholine have been demonstrated t o occur between the bathing medium and the intact central nervous tissues of P. americana (Treheme and Smith, 1965a; Eldefrawi and O’Brien, 1967). Despite the rapid influx of labelled acetylcholine molecules measured in these investigations the calculated extracellular acetylcholine concentration was found to be 8.1 x M in preparations bathed with 1O-* M acetylcholine (Treherne and Smith, 1965b). In these exReriments the products of the hydrolysis of acetyL3Hcholine accumulated largely as acetate and glutamate with the nerve cord tissues. It appeared, from these results, that an appreciable hydrolysis of acetylcholine occurred towards the periphery of the

INSECT BLOOD-BRAIN BARRIER

299

nerve cord. Thus, for example, when whole nerve cords were bathed in a solution of radioactive 10-2 M acetylcholine the concentration of this substance within the nervous tissues showed only a slow approach to the level obtained after a one second exposure in the presence of eserine. The amount of radioactivity taken up during this brief exposure of labelled acetylcholine solution was found to be roughly equivalent to that contained in the nerve sheath. It was postulated, therefore, that a considerable hydrolysis of the applied acetylcholine must have occurred at the periphery of the ganglion, possibly within the nerve sheath itself (Treherne and Smith, I965b). The peripheral sites of cholinesterase activity, which were shown to be localized on the glial membranes bordering peripheral extracellular channels (Smith and Treherne, 1965; Treherne and Smith, 1965b), would be strategically placed to effect hydrolysis of acetylcholine penetrating in to the perineurium and neural lamella. It would appear, therefore, that the access of acetylcholine molecules to the synaptic surfaces may be limited not only by diffusion barriers, but also by the presence of a “metabolic barrier” situated at the periphery of the ganglion. VI. CONCLUDING REMARKS

The overall picture which emerges from the measured electrophysiological responses of nerve cells is of a restricted access of ions and molecules to the extraneuronal fluid. In this sense it can be clearly stated that insects possess a well developed blood-brain barrier system; our present concept being that the main restriction to intercellular diffusion of water-soluble substances to the neuronal surfaces from the blood, results from the presence of intercellular occlusions at the inner ends of the perineurial clefts. It is also clear from radioactive studies that, despite this peripheral restriction to intercellular ion movements, relatively rapid steady-state exchanges nevertheless occur between the blood and the central nervous system. This is well exemplified by the rapidly-exchanging 22Na fraction (which, according to Tucker and Pichon (1972b) amounts to 45% of the exchangeable sodium) observed under conditions of maximal peripheral restriction to intercellular ion movements postulated on the basis of the observed extraneuronal potential changes in the cockroach nerve end. The steady-state exchanges of radioactive ions and molecules observed under these conditions strongly suggest the possibility that

300

J . E. TREHERNE AND Y. PICHON

rapid movements occur via an intracellular pathway within the nervous system. The most obvious candidate for this would seem to be the glial cytoplasm. This concept would accord with our present knowledge of the ultrastructural organization of insect central nervous tissues. In particular the apparent existence of low-resistance pathways linking the perineurial and underlying glial cytoplasm would ensure the rapid movement of substances within the glial compartment. The uptake of substances through the outwardly facing perineurial surface would be facilitated by the large effective membrane area resulting from the extensive infoldings. Phenomena such as exchange-diffusion mechanisms, for sodium at least, and active transport would also explain the rapid and extensive steady-state exchanges observed in intact central nervous preparations. It should, in addition, be borne in mind when considering the electrophysiological results in relation t o those obtained using radioisotopes that in the former case we are studying the effects of net ion movements resulting from substantial changes in concentration in the bathing medium. In these conditions electro-chemical gradients are created which may contribute to the modification of permeability characteristics of the system as compared with those pertaining in the steady state conditions. These factors are represented diagrammatically in Fig. 22. The evidence presented above, showing a significant restriction to diffusion of substances between the blood and the fluid bathing the neuronal surfaces in insects, contrasts markedly with the situation in the only other arthropod species which has been investigated: the shore crab, Curcinus maenas (Abbott, 1969, 1970). In this crustacean there appears to be no equivalent restriction to ionic and molecular exchanges with the blood which in this species is channelled into the brain by a well-developed capillary blood supply (see also Sandeman, 1967). Furthermore, as was emphasized in a recent review (Treherne and Moreton, 1970), there is no substantial evidence for the existence of noticeable restriction in access to the neuronal surfaces in the other higher invertebrates which have been studied. The closest parallel to the blood brain barrier of insect species appears to be that encountered in higher vertebrate animals. In mammalian brain, as in insect central nervous tissues, the restricted access to the fluid surrounding neurones appears to result from the presence of intercellular occlusions interposed between the blood and the extraneuronal fluid. These occlusions are in the form of “tight” junctions between the lateral membranes of the capillary

4 I

+t

fR2-0

..... ................................................... (7)

...I......

/

Fig. 22. Simplified and speculative diagram of the exchange properties of the nervous tissue in Penplaneta americanu L. The large circles 1, 2 and 3 represent respectively the presumed extracellular, glial and axonal compartments. The continuous lines indicate the extracellular pathway, the interrupted lines an intracellular pathway. The resistors represent the proposed main sites of restriction. R, = axonal membrane resistance; R3+: inner glial mesaxon membrane resistance; R+,: restriction in the mesaxon channels; R2-l: glial membrane resistance; R2-; outer perineurial membrane resistance; Rl-o: tight junctions. Rl-o and R2-o are not readily distinguished by physiological experiments and are likely to be coupled (as indicated by dotted line). The arrows indicate the possible pathways used by aa Na ions to leave the nervous tissue in radio-isotope experiments. In fully intact preparations, the effective value of Rl-o is large compared to R,-o (because of the large membrane wea of the perineurial cells). The overall resistance of the outer barrier (R) approaches Rz-o and the permeability of the system to different molecules is a function of the properties of the outer perineurial membrane. In stretched preparations Rl-o is reduced and possibly Rz-o . The value of R is lower than in intact conditions: the nerve sheath is less effective in regulating the movements of ions and molecules from and into the nervous system. In very stretched or desheathed preparations, R approaches or equals zero and the restriction to the movements of ions and molecules from and into the axons is a function of the values of R,, R3-, and R3-2 . In non-steady state conditions, electrochemical gradients are created which affect the movements of ions through the system. For example, in high external K+,a potential is created between the outer and the inner perineurial membranes (the extraneuronal potential) which opposes inward movements of K + ions; this effect is more or less short circuited by R,-,, in mechanically treated preparations. In Na+ free solutions, R2-,, which is large in resting conditions (due to the properties of the outer perineurial membrane) becomes even larger due to the building-up of a potential across the perineurium (possibly associated with the disappearance of exchangediffusion movements of sodium); this restriction is also more or less short-circuited by Rle0 in mechanically treated preparations. (Modified from Tucker and Pichon, 1972b.)

302

J. E. TREHERNE AND Y . PICHON

endothelium (in mouse, hamster and Necturus) or the perivascular glial end-feet (in elasmobranchs) which, as in their insect counterparts, restrict the entry of peroxidase (Brightman and Reese, 1970). Another striking similarity between the vertebrate and the insect situations are the presence of appreciable potential changes which have been recorded between the cerebrospinal fluid and the blood following alterations in the potassium concentration of the blood (c.f. Cameron and Kleeman, 1970). It would seem reasonable to conclude that these potential changes might have a functionally similar basis to those proposed (p. 285) for the insect central nervous system. A diagrammatic representation of our current conception of the blood-brain barrier in insects related to those for vertebrates, a crustacean and an annelid is given in Fig. 23. A distinctive feature of the physiological organization of insect central nervous tissues is their ability to maintain ionic concentrations in the extra-axonal fluid which differ markedly from those of the haemolymph. This is seen most dramatically in phytophagous species, such as C. morosus and M. sexta, in which the sodium concentration in the fluid bathing the axon surfaces may be approximately an order greater than that of the haemolymph. This phenomenon may not, however, be confined to phytophagous insects, for as has been pointed out, some electrophysiological evidence can be interpreted on the assumption that the extra-axonal sodium concentration of P. americana (which has a relatively high haemolymph sodium level) exceeds that of the haemolymph by a factor of approximately 2.3. The precise nature of physiological mechanisms involved in extra-axonal sodium regulation still remains to be elucidated. It is likely that relatively high extracellular sodium concentrations could be maintained by the anion groups associated with the acid mucopolysaccharide (c.f. Treherne, 1962a). However, it seems clear that the thermodynamic activity of sodium ions in such a fixed-anion system would only be equivalent to that in the haemolymph or bathing medium and could not, therefore, contribute to an elevation of the effective sodium concentration of the extra-axonal fluid (Treherne and Maddrell, 1967b; Treherne, 1967). It seems reasonable to suppose, however, that any sodium ions associated with extracellular fixed-anion groups would tend to act as a cation reservoir which might function in the short-term maintenance of the ‘extra-axonal sodium level. It could thus be envisaged that any fluctuations in the sodium concentrations in the vicinity of the


303

Leech blood

nl

blood

blood

.

Insects

Teleost

. . ..

, ........, ...... . . . ... ,:.:.:.:.'.'.'.'.'*.'.'... . . . . ....:.:. ::::.:.: .:.:.:.: :.:.:.:. .'.'.:.:

blood

Crustacean

blood

Elosrnobronchs

Fig. 23. Diagrammatic representation illustrating some current concepts of the structural basis of the insect and vertebrate blood-brain barriers, in comparison with the situation in an annelid and a crustacean species. The interpretation of the situation in vertebrate animals is based on data summarized by Brightman and Reese (1970), that for Curcinus maenus from Abbott (1969, 1970) and for Hirudo medicinalis from Coggeshall and Fawcett (1964), Nicholls and Kuffler (1964) and Kuffler and Potter (1964). The representation of the situation in the insect species is based on the various data presented in this article, especially that of Lane andTreherne (1969,1970,1971) and Lane (1972.)

a: axon; bm: basement membrane; e: endothelium; fbs: fat body sheath (present in some species); g: glia; nl: neural lamella; p: perineurium.

304


axons, resulting from their electrical activity, would be compensated for by a release of cations associated with the fixed-anion system. The only other hypothetical physiological mechanism which has been advanced attempts to relate extra-axonal sodium regulation to the activity of glial elements (Treherne, 1967; Treherne and Maddrell, 1967b; Pichon and Boistel, 1967; Pichon 1969a). It has further been tentatively suggested that the positive extracellular potentials demonstrated by Pichon and Boistel (1967) might be a manifestation of a glial-mediated electrogenic sodium pump which could be involved in the maintenance of the high concentration of sodium ions in the fluid bathing the axon surfaces relative to that of the haemolymph (Treherne and Moreton, 1970). This hypothetical mechanism would also accord with the concept, advanced above, of an intracellular low-resistance pathway linking the perineurium with the innermost glial membrane adjacent to the axon surfaces in P. americana. Such an intracellular route would form an obvious pathway for the movement of sodium ions to the glial cytoplasm immediately adjacent to the extra-axonal fluid. It should be borne in mind considering the validity of the provisional hypothesis, outlined above, that the positive extracellular potential could arise from mechanisms other than glial electrogenic sodium pumps. It is conceivable, for example, that the positive extracellular potential could result from the juxtaposition of the positively-charged outer neuronal surfaces with the extracellular negatively-charged acid mucopolysaccharide in a manner analogous to that postulated for vertebrate malignant trophoblastic cells (Hause et al., 1970). However, the provisional hypothesis advanced above represents a system which should be susceptible to experimental analysis and, for this reason, may be of assistance in future research in this area of neurophysiology. Whatever the nature of the extra-axonal sodium regulating system in insects it seems obvious that the restriction to intercellular diffusion offered by the perineurial “tight” junctions would greatly facilitate the functioning of the system by reducing the escape of sodium ions from the extracellular system. The perineurial restriction to the intercellular diffusion of watersoluble ions from the haemolymph or bathing medium appears to exist, as was described above, in parallel with relatively large rapidly-exchanging ion fractions in the central nervous system of P. americana. These rapidly-exchanging ion and molecular components, demonstrated in radioisotope experiments, have been tentatively


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identified as being of glial origin. The existence of such rapidlyexchanging glial fractions suggest a possible mechanism by which nutrient substances could be transported from the haemolymph to the nerve cells in the presence of the peripheral intercellular diffusion barrier. The rate of uptake of nutrient molecules from the haemolymph is presumably enhanced by the relatively large outwardly-directed perineurial surface resulting from the extended and tortuous perineurial clefts: the rapid transport through the glial cytoplasm being facilitated by the low resistance junctions between adjacent glial cells. The existence of such a glial transport system was foreshadowed by the histochemical observations of Wigglesworth ( 1960) who demonstrated a sequential deposition of glycogen, through the nervous systems of starved cockroaches following injections of glucose into the blood. The sequence was terminated by the transfer of material from the glial cytoplasm to neuronal cell body via the trophospongium of Holmgren. It is possible, therefore, to visualize a dual system of exchanges in the insect central nervous system : a restricted access of water-soluble ions and molecules into the extracellular system, which is paralleled by relatively rapid exchanges with the glial cytoplasm. As should be apparent from the above account this arrangement facilitates both the regulation of the ionic environment of the neurones and, despite the peripheral restrictions to intercellular diffusion, allows rapid and effective exchanges of nutrient substances to occur between the haemolymph and the neurones.

ACKNOWLEDGEMENTS

We are indebted to Dr. N. J. Lane and Miss Y. Carter for their help in preparing some of the electron micrographs and to Mr. J. Rodford for supplying some of the illustrations included in this article.

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NOTE ADDED IN PROOF Since this article was submitted for publication the following relevant observations have been reported. A recent study, employing en bloc uranyl acetate staining (Karnovsky, 1967), has revealed that the perineurial intercellular occlusions in P. americana consist of septate desmosomes, gap junctions and tight junctions (zonula occludens) (Lane and Treherne, 1972). Microperoxidase (h4W 1900) showed extensive penetration of intercellular clefts and limited penetration of septate desmosomes. There was no evidence of penetration into the underlying extracellular spaces. Colloidal lanthanum was seen t o penetrate both septate desmosomes and gap junctions. It is concluded, therefore, that its relatively limited penetration to the underlying extracellular system probably results from its restricted access through the tight junctions (see p. 268). An account has been published on the efflux of 2%a from nerve cords of Carausius morosm and Periplaneta americana (Weidler e t al., 1971). Isolated nerve cords were used in these experiments and were “loaded” in radioactive solutions for periods of up to 14 h. The authors extracted no less than six components from their desaturation curves which were obtained using efflux periods of 26 h. With shorter periods, of about lOmin, only two efflux


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components were recognized. These results, which the authors claim to be consistent with the previous electrophysiological findings (see p. 28 l), appear to have little physiological significance. For example, the long duration of these experiments (up to 40 h) was quite inappropriate, especially as it is known that isolated nerve cords show significant changes in Na' and K + concentrations after only 1.5 h in normal physiological solution (Tucker, unpublished observations). Furthermore, no statistical analyses were employed in this investigation, neither was any attempt made either to correlate these components with any functional properties of the system or to test the excitability of the isolated nerve cords during the long experimental periods employed. The hypothesis that a significant proportion of the observed radioactive flux in short duration experiments originates in the fat body is also unacceptable for similar efflux components have been observed from isolated connectives of Periplaneta (see p. 294) which were free of fat body deposits (Tucker and Pichon, 1972b).

REFERENCES Karnovsky, M. J. (1 967). The ultrastructural basis of capillary permeability studied with peroxidase as a tracer. J. Cell Biol. 3 5 , 21 3-236. Lane, N. J. and Treherne, J. E. (1972). Studies on perineurial junctional complexes and the sites of uptake of microperoxidase and lanthanum by the cockroach central nervous system. Tissue and Cell (In press). Weidler, D. J., Myers, G . C., Gardner, P. J., Bennet, A. L. and Earle, A. M. (1971). Defects in the experimental design of radioisotope studies on the insect nerve cord. 2.vergl. Physiol. 7 5 , 352-366.


Insect Sperm Cells BACCIO BACCETTI Institute of Zoology. University of Siena. Italy

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VII .

VIII .

Introduction . . . . . . . . . . . . . . . . . The Cell Surface . . . . . . . . . . . . . . . . The Acrosomal Complex . . . . . . . . . . . . . A. The Typical Triple-layered Insect Acrosomal Complex . B. The Bilayered Acrosomal Complex. . . . . . . . . C. The Acrosomal Complex with Only Two Outer Layers . D. Monolayered Acrosomal Complex . . . . . . . . E. Total Absence of Acrosomal Complex . . . . . . . The Nucleus . . . . . . . . . . . . . . . . . A . Nuclear Shape . . . . . . . . . . . . . . . B. Submicroscopic Structure . . . . . . . . . . . C . Chemical Characteristics . . . . . . . . . . . D. Physical Characteristics . . . . . . . . . . . . The Centriolar Region . . . . . . . . . . . . . . A . The Centriole . . . . . . . . . . . . . . . B. The Centriole Adjunct . . . . . . . . . . . . C. The Initial Segment of the Axoneme . . . . . . . . The Axial Flagellar Filament or Axoneme . . . . . . . A. The Microtubules . . . . . . . . . . . . . B. The Central Sheath . . . . . . . . . . . . . C. The Link-heads . . . . . . . . . . . . . . D. The Coarse Fibres . . . . . . . . . . . . . E . The Axonemal Matrix . . . . . . . . . . . . Mitochondria . . . . . . . . . . . . . . . . . A. Normal Mitochondria . . . . . . . . . . . . B. Mitochondria Transformed into Derivatives with a Crystalline core . . . . . . . . . . . . . . . . . C. Absence of Mitochondria . . . . . . . . . . . Accessory Ordered Flagellar Bodies . . . . . . . . . . A . Structured Bodies Flanking Normal Mitochondria . . . B. Structured Bodies Flanking the Mitochondria1 Derivatives with a Crystalline Matrix . . . . . . . . . . . C. Structured Bodies Replacing Mitochondria . . . . . 315

316 317 324 324 326 327 321 328 328 328 329 331 331 332 332 333 335 336 338 349 349 350 352 354 354 354 360 363 363 365 365

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Spermatozoa Possessing a Double Flagellar Apparatus or Being Devoid of it. . . . . . . . . . . . . . . . . . A. The Paired Spermatozoa . . . . . . . . . . . B. SpermatozoaPossessing Two Axonemes . . . . . . C. Non-flagellate Spermatozoa . . . . . . . . . . X. Motility. . . . . . . . . . . . . . . . . . . A. Motile Mechanisms . . . . . . . . . . . . . B. Metabolic Aspects of Motion . . . . . . . . . . C. The Problem of Sperm Capacitation . . . . . . . XI. SpermatozoaPolymorphism and Genetics . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . IX.

367 367 369 370 374 374 380 38 1 382 384 384

I. INTRODUCTION

Among the animal phyla that have adapted themselves permanently to land life, the spermatozoon had to undergo thorough changes in its structure, metabolism and motility as a consequence of the onset of internal fertilization. The aquatic environment having been abandoned, spermatozoa were required to swim in a completely different medium, which was produced usually by the two partners at fertilization. Annelida, Mollusca, Arthropoda and the vertebrates have separately, but sometimes with amazing analogies, devised new forms of male gametes. In some arthropod groups still breeding in water, a “primitive” sperm model is retained which might be referred to as “aquatic” in type, e.g. in the Merostomata (AndrC, 1965; Baccetti, 1970; Shoger and Brown, 1970) and in the crustaceans Mystacocarida (Brown and Metz, 1967), Cirripedia and Branchiura (Brown, 1966, 1970). The outstanding features of this “aquatic” sperm type can be found in the most ancient representatives of almost all phyla, including Amphioxus among the Chordata (Baccetti et al., 1972f). These “primitive” features are represented by a roundish head, a great number of normal mitochondria rich in cristae and a relatively short flagellum with a plain axial filament simply organized according to the 9 + 2 pattern. This model was inherited by all arthropod classes, including those which are now definitely terrestrial. If allowance is made for some slight variations, this model is retained by primitive Arachnida, the scorpions (AndrC, 1963); among primitive Myriapoda, by the Pauropoda, Symphyla (Rosati et al., 1970) and Chilopoda (Horstmann, 1968); as well as by the Collembola and Diplura among primitive insects (Dallai, 1967, 1970; Baccetti et al., 1972d).

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When we follow the pathway of their evolution as revealed by extant forms, it may be observed that the Arachnida, Myriapoda and Insecta have subsequently refined their spermatozoa. Most Arachnida and Myriapoda produce “encysted” sperms (Baccetti, 1970) which are subsequently released in the female genital tract. All the spiders exhibit a peculiar 9 + 3 flagellum (Baccetti et al., 1970d; Reger, 1970) which is apparently of little functional significance because the encysted sperm is transported passively. As a consequence, the most evolved of the two lines, Opilionids (Reger, 1969) and Acarina (Reger, 1963; Breucker and Horstmann, 1968) on one side, and high Diplopoda (Reger and Cooper, 1968; Horstmann and Breucker, 1969a, b; Reger, 1971) on the other, have non-flagellate spermatozoa. Conversely, the spermatozoon of insects has evolved in the direction of increased motility: its shape, structure, metabolism and locomotory capabilities are tremendously differentiated. In many instances they attain an extreme specialization which is related to: (1 ) the mode of sperm transmission (internal fertilization always being the case); (2) the duration of its survival in the male genital tract or within spermatophores and spermathecas; (3) the type of fluid they have to swim in; and (4) the complexity of the egg envelopes they must penetrate. Typical features of the insect spermatozoon (Fig. 4) are the following: a generally very slim shape with an extremely elongated head; a bi- or three-layered acrosomal complex; an exceedingly long tail whose axial filament is flanked by accessory structures usually derived from mitochondria1 transformations. Along these lines evolutionary changes have resulted in a highly diversified and, in some respects, puzzling picture. Since some insight into the structure and function of spermatozoa has been gained during the past few years, it seems worth while to determine how far our knowledge on this subject has progressed. 11. THE CELL SURFACE

.

Mature insect spermatozoa are surrounded by the typical triple-layered membrane which appears markedly asymmetrical in electron microscopical sections (Fig. 9) due to the fact that with all fixation and staining procedures its outer electron opaque layer consistently is as thick as the intermediate light (30 A ) and opaque

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inner (20 A) layers put together. According to Baccetti et al. (1 97 la) both opaque layers show acid phosphatase activity (Fig. 1C). They appear smooth after freeze-etching whereas the intermediate transparent layer contains a number of scattered globules (Fig. IA). The outer layer reacts positively to all the methods for glycoproteins, which suggests that this membrane resembles a common plasma membrane with a strongly thickened glycocalyx. This model in which, moving from the inside to the outside, the layers are consecutively some 20, 30 and 60 A in thickness and the glycocalyx appears structureless, is shared by the vast majority of insect spermatozoa. It was referred to as the “fruit-fly type membrane” by Baccetti et al. (1 97 1a). In addition, three progressively more elaborate arrangements are known. The first one (Fig. 1) was described in Aphaniptera and given the name of “flea type membrane” again by Baccetti et al. (1971a). In this instance, the outer sperm wall is about 200 A thick, since a 130 A thick glycocalyx is attached to the normal plasma membrane which averages 70 A . The glycocalyx has a peculiar structure. It is arranged around the spermatozoon in transverse striae, with a periodicity of some 140 A , overlying the outer surface of the plasma membrane (Fig. IB). A more complex arrangement (Fig. 2) was found among locusts (“locust type membrane”) by Baccetti et al. (1971a). Here, as was described in earlier observations (Roth, 1957; Kessel, 1967), the sperm wall is as thick as 400 A, due to a glycocalyx exceeding 300 A in thickness (Fig. 2A) and comparable to the fuzzy coat of many protozoa. This coat is made up of rodlets 300 A long and 70 A wide inserted almost perpendicularly into the sperm surface. They are arranged in tetrads in which every rodlet is 2 0 A apart from its neighbour (Fig. 2B). Each tetrad is about 70 A away from the others. Both the striated envelope of the Aphaniptera spermatozoon and the fuzzy one in locusts are derived from an abundant amorphous glycoprotein material coating the spermatid. This material becomes organized over the plasma membrane during sperm maturation. More elaborate and less obvious is the origin of the most complex type of sheath so far known, namely that of Lepidoptera, which was described in butterflies by AndrC (1 959, 1962) and Phillips (1 970b), and in moths by Riemann (1970). This type, however, has been recently confined to the eupyrene sperms (Phillips, 197 1). This sheath (Fig. 3A) is made up of numerous appendages, called lacinate

Fig. 1. The cell periphery of the spermatozoon of Ctenocephalus canis. (From Baccetti et aZ., 1971a.) (A) Frozen-etched preparation, showing the striation of the outer surface (s) and the granulated (g) space of the cell membrane. ~60,000.(B) Longitudinal section of the plasma membrane (p) and of the array of fibres striating the cell coat (s). ~150,000.(C) Acid phosphatase on plasma membrane (p) and on the outer cell coats (s). ~80,000.

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Fig. 2. The cell periphery in the spermatozoon of Aiolopus strepens. (From Baccetti et al., 1971a.) (A) Cross section of mature sperms, showing the brush-structured outer coat (c) surrounding the plasma membrane (p). ~120,000.(B) Negative staining of the fragmented cell membrane, showing the frontal view of the rodlets arranged in tetrads (s). ~360,000.

Fig. 3. The lacinate coat of the Lepidopteran spermatozoon. (From Phillips, 1971.) (A) Transverse section of eupyrene testicular spermatozoa showing the radiate outer appendages. x 120,000. (B) Transverse section of eupyrene spermatozoa after entering the female. The radial appendages are substituted by two layers of extracellular material (arrows). x 106,000.

Fig. 3 AIP-14

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appendages by Andre (1959, 1962), radially arranged all around the spermatozoon. Each appendage consists of many juxtaposed laminae 90 A apart from one another. One of these appendages is lattice-like instead of laminar and lies at the level of doublet 1 of the flagellum. This crown of appendages seems to be derived from a peculiar amorphous small body referred to as “light band” by Yasuzumi and Oura (1964). In the spermatids, this body arises externally and differentiates into the radially arranged laminar structures. The glycocalyx is not the only structure to grow and develop during the last phases of spermiogenesis. There are also striking changes in the plasma membrane associated with the change in cell shape which occurs when the roundish spermatid turns into an impressively elongated spermatozoon. In the mature spermatid, progressively more complex membrane systems develop. These are flat cisternae which envelope the main organelles, notably the axial filament and mitochondria, demarcating the form of the future sperm and excluding wide regions of the spermatid cytoplasm which will later be shed. These membranes were regarded as endoplasmic reticulum by Yasuzumi et al. (1958) as well as by Ito (1 960). It was pointed out by Baccetti et al. (1 972b) that they are derived from the Golgi complex, and that during sperm maturation they usually fuse with the limiting membrane. In some places they simply adhere to the inner surface of the plasma membrane, doubling its thickness. Elsewhere a total fusion takes place, eventually resulting in a simple plasma membrane; again at other sites the original plasma membrane is eliminated and replaced by a newly formed membrane of Golgi origin. The morphological significance of these membranes, which clearly originate from the Golgi complex, is still debated. In spermatocytes, Sakai and Shigenaga (1 967) favour the view that the tubular endoplasmic reticulum originates from the Golgi. It may perhaps be preferable (Baccetti et al., 1972b) to call them Golgiderived membranes, mindful of the fact that many of them are going to build the definitive limiting membrane. Alteration of the membranes does not stop at the end of spermatogenesis, but undergoes further modifications during sperm transfer from the male to the female. The elaborate lacinate appendages of the eupyrene spermatozoa in the Lepidoptera undergo breakdown in the male genital duct. The material they are composed of is rearranged into concentric bands encircling the formerly naked apyrene sperms (Phillips, 1971). Once they have reached the female (Fig. 3B), both eupyrene and apyrene sperms possess identical

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Fig. 4. Schematic drawing of conventional models of a mature insect spermatozoon. The acrosome region is the three-layered one of Aiolopus (Orthoptera), the centriole region the radiated one of Bacillus (Phasmoidea), the tail the most complete known, of Tenebrio (Coleoptera): A, acrosome; AB, accessory bodies; AC, centriole adjunct; AT, accessory tubules; CC, central cylinder; CF, coarse fibres; CS, central sheath; CT, central tubules; D, doublets; DD, dissociated doublets; IC, inner cone; EL, extraacrosomal layer; GM, Golgi derived membranes; LH, link-heads; MD, mitochondria1 derivatives; N, nucleus: OC, outer cylinder; PM, plasma membrane; RL, radial links; RLA, radial laminae.

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envelopes. In moths, these envelopes differentiate into two parts having different paracrystalline structures (Riemann and Thorson, 1971). On the contrary, the “fuzzy coat” of the locust sperm is completely digested in the female genital tract (Renieri and Vegni, 1972). 111. THE ACROSOMAL COMPLEX

A multi-layered acrosomal complex is present among the most primitive insects such as the Diplura, Collembola and Thysanura (Dallai, 1970, 1972; Baccetti et al., 1972d). There are but small increases in complexity of this basic arrangement up to the most evolved Pterygota, like the Coleoptera (Baccetti et al., 1972a) and Hymenoptera (Hoage and Kessel, 1968). For these latter examples, therefore, it is possible to propose a general model of acrosomal structure for the whole class. The various modifications exhibited by single orders and species can be derived from this model and always consist of various structural simplifications. In extreme cases, this organelle is completely absent. The insect acrosome, as is the rule for all animals, is typically derived from the Golgi complex (the earliest evidence in this regard was provided by Beams et al. (1 956), Clayton et al. (1 958), Ito (1960), Gatenby and Tahmisian (1 959) and others). A. THE TYPICAL TRIPLE-LAYERED INSECT ACROSOMAL COMPLEX

This arrangement, with few variations, is present in almost all the Pterygota. It has been described in great detail for Orthoptera and Blattoidea (Kaye, 1962; Eddleman et al., 1970; Baccetti et al., 1971c), in Mecoptera (Baccetti et al., 1969c), in Aphaniptera (Baccetti, 1968), in the coleopteran Tenebrio (Baccetti et al., 1972a) and in the honey-bee (Hoage and Kessel, 1968). Generally, three juxtaposed layers (sometimes described as concentric) are encountered on the inside of the plasma membrane (Fig. 5A). The outermost “extraacrosomal layer” is an aggregation of granular cytoplasmic material which is concentrated between the plasma membrane and the acrosome proper during spermiogenesis. It occurs in a€l the instances mentioned here, but is particularly abundant in the more primitive orthopterans (Kaye, 1962; Shay and Biesele, 1968; Baccetti et al., 1971c). Its significance is utterly obscure. Beneath it the acrosome proper is found. This layer originates from the proacrosomal granule of the spermatid which is synthesized by

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Fig. 5. Three different arrangements of the acrosome complex in insects. (A) The three-layered model of Tenebrio: a, acrosome; el, extraacrosomal layer; ic, inner cone; pm, plasma membrane; n, nucleus. ~60,000. (From Baccetti el al., 1972a.) (B) The bilayered model of Campodea: a, acrosome; ic, inner cone. ~48,000.(From Baccetti and Dallai, 1972.) (C) The monolayered model of Chloeon: a, acrosome; n, nucleus. ~60,000 (From Baccetti et aZ., 1969b.)

the Golgi apparatus. Synthesis is most frequently associated with the concave side of its cisternae (Gatenby and Tahmisian, 1959; Kaye, 1962; Phillips, 1966a). In a single case it was seen to arise from the convex side (Phillips, 1970b). Histochemical investigations (Zylberberg, 1969; Baccetti et al., 1971c) have shown that the acrosome consists of a rather stable glycoprotein which is resistant to extraction. By negative staining the acrosome reveals a 95 vertical period and a 25 A horizontal period (Warner, 1971). The acrosome is

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entirely surrounded by its own triple-layered membrane, which previously encircled the proacrosomal granules derived from the Golgi (Kaye, 1962), and contains acid phosphatase (Baccetti et al., 1972b). Within the acrosome proper the third layer is encountered. This may be rodlike or conical, in the latter case being called “inner cone”. This is a compact structure, which is formed in the spermatid in an interspace between the acrosomal granule and the nuclear membrane, independently of the acrosome proper or of the Golgi region (Kaye, 1962). The characteristic interstitial membrane, which in the spermatid forms between the nuclear membrane and the proacrosomal granule (Kaye, 1962), is seen to break down at the site where this inner rodlet arises. Generally, no trace of the interstitial membrane is left in the mature sperm. The inner rodlet is endowed with histochemical properties diverging considerably from those of the acrosome; it consists of a protein core surrounded by a glycoprotein halo (Baccetti et al., 197 1 c). Information as to its origin and significance is entirely lacking. According to its position and ontogenesis, it might be regarded as comparable to the vertebrate perforatorium, which also contains carbohydrates (Sandoz, 1970), but its chemical nature is unknown. So far, the functional interpretation of the different components of the insect acrosomal complex is difficult to determine since even the mammalian one is as yet poorly understood. In mammals, according to Austin (1948) and Wada et al. (19S6), the acrosome contains hyaluronidase, while the perforatorium is believed to contain lysine (see Dan, 1967). Among the insects, the acrosomal hyaluronidase is completely absent (Baccetti et al., 197 1c). However it is not known whether lysines are similarly concentrated in the acrosome. This triple-layered acrosomal arrangement is generally conical in shape. However, in the tettigonioid Orthoptera it is arrow-like, and its stratification is also detectable inside the arrow limbs (Phillips, 1970b; Baccetti et al., 1 9 7 1 ~ ) In . the spermatozoon it generally lies apically to the nucleus. B. THE BILAYERED ACROSOMAL COMPLEX

This arrangement is very similar to the triple-layered acrosomal complex described above, the only difference is the absence of the outer extraacrosomal layer (Fig. SB). This arrangement is typical of the Apterygota. The extraacrosomal layer is consistently lacking in

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Collembola (Dallai, 1970), Protura, Diplura (Baccetti et al., 1972d) and Thysanura (Dallai, 1972). In this respect the bilayered arrangement might be regarded as more primitive than the triple-layered one, rather than the reverse. But, pending clarification of the nature and function of the extraacrosomal layer, nothing can be said about it at present.

C. THE ACROSOMAL COMPLEX WITH ONLY TWO OUTER LAYERS

This is a modification of the triple-layer arrangement; it consists of the acrosome proper and the extraacrosomal layer. Examples of this arrangement are found in phasmoidea (Baccetti et al., 1972b) and among a number of Homoptera (Folliot and Maillet, 1970). When the acrosome of Hemiptera has reached a considerable length, it becomes enriched by inner and outer microtubules which maintain its rigidity (Payne, 1966; Tandler and Moriber, 1966; Folliot and Maillet, 1970) and are still present in the mature spermatozoon. Since the inner rodlet is found in the more primitive arthropods and in the apterygotes as well, this arrangement might be interpreted as an involution of the triple-layered one. In Bacillus, glucose-6phosphatase is detectable in a restricted basal zone (Baccetti et al., 1972b) which might indicate a specialized area, although any speculation in this regard is far from easy.

D. MONOLAYERED ACROSOMAL COMPLEX

This structure consists of the acrosome alone (Fig. SC), enveloped by the acrosomal membrane and the plasma membrane. It has been reported in many groups: it occurs regularly in Ephemeroptera (Baccetti et al., 1969b), in Plecoptera (Baccetti et al., 1970e), in many Heteroptera (Barker and Riess, 1966; Herold and Munz, 1967; Mazzini, 1970), in Corrodentia , Mallophaga and Thysanoptera (Baccetti et al., 1969d), in those Trychoptera which possess the acrosome (Phillips, 1970b), in Lepidoptera (Phillips, 1971), in Diptera (Bairati and Perotti, 1970; Perotti, 1969; Tates, 1971, in Drosophila; Phillips, 1966b in Sciara) and in the Psychodidae (Baccetti et al., 1972c). This type of acrosome is always of very small size, and is often displaced to a lateral position with respect to the nucleus.

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E. TOTAL ABSENCE OF THE ACROSOMAL COMPLEX

This is occasionally found in some orders in which certain species possess an acrosome. Some instances are found in some of the more highly evolved species of the Isoptera (Baccetti et al., 1972e); in Coccoidea among Rhynchota, (Robison, 1966); in some species among Trychoptera (Baccetti et al., 1970e; Phillips, 1970b), in Oryctes among Coleoptera (Furieri, 1963a). In a whole order, Neuroptera (Baccetti et al., 1969c), the acrosome seems to be consistently lacking. This condition seems also due to involution. All the information summarized above seems to suggest that the triple-layered arrangement, typical of the pterygotes, was derived from a primitive bilayered acrosome model (rodlet and acrosome sensu stricto) as still retained by Apterygota. In some of these forms the acrosome may lose its rodlet and become bilayered (but the two outer layers are retained, hence not corresponding to the two found in Apterygota), monolayered or disappear altogether. The acrosomal layer itself is the only one present whatever the arrangement. IV. THE NUCLEUS

In a recent review on the sperm nucleus, Chevailler (1 970) points out that it is usually given less attention than the other sperm organelles, possibly due to its structural homogeneity and the uniformity of its transformations during spermiogenesis in various species. However, the survey carried out by this author brings into focus many important features, which will be discussed in the following section. A. NUCLEAR SHAPE

Among the insects, the sperm nucleus is as a rule fairly elongated, spindle-shaped anteriorly and truncated posteriorly. It is considerably compact, occasionally helicoidally arranged (Fig. 6b), sometimes even coiling around the flagellum as in the Mecoptera or Thysanura (Baccetti et al., 1969c, d). In Dahlbominus (Hymenoptera) some sperms have the helix twisted to the right, others to the left (Wilkes and Lee, 1965). Only in the spermatozoa provided with two axial filaments does the nucleus display a short flat shape as found in Anoplura and Mallophaga (Baccetti et al., 1969d). Aberrant

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non-flagellate sperms found in the most highly evolved Isoptera have a spheroidal or flattened nucleus (Baccetti e t al., 1972e). B. SUBMICROSCOPIC STRUCTURE

As a rule, in the mature insect sperm the nucleus appears compact and homogeneous in electron micrographs. This structure is attained during spermiogenesis. The chromatin in the young spermatid thick (Yasuzumi and Ishida, 1957; contains fibrils some 30-40 Gibbons and Bradfield, 1957; Dass and Ris, 1958; Gall and Bjork, 1958; Nebel, 1957), which are rapidly converted into thicker fibrils (1 00-200 arranged longitudinally (Werner, 1966; Chevaillier, 1970). These fibres then fuse into laminae which sooner or later coalesce. In the Apterygota the laminar pattern is also retained by the nearly mature spermatozoon (Bawa, 1964; Dallai, 1967) and the nucleus becomes compact just before its expulsion. In some instances, the nucleus instead of being homogeneous, exhibits a honey-comb texture (Yasuzumi and Ishida, 1957). The nuclear material does not undergo uniform morphological changes during spermiogenesis. Sometimes the central zone is the first to condense, at other times it is the periphery (Chevaillier, 1970). Among Psyllidae (Le Menn, 1966) only half the nucleus is occupied by chromatin, even in the mature sperm. During spermiogenesis the nuclear envelope is surrounded by a layer of microtubules which function in the compression and elongation of the nucleus, and which will disappear in the mature sperm (see, in particular, Kessel (1966, 1967)). DNA is seen to aggregate preferentially at the microtubule level (Baccetti e t al., 1972b) as is the case in most animals (Ferraguti and Lanzavecchia, 1971). Wide portions of the nuclear membrane are seen to form blebs which are pinched off into vesicles and dispersed into the cytoplasm (Kessel, 1970). Pores progressively decrease in number until they are no longer evident in the mature sperm. In some instances, however, the nucleus seems to take part in the elaboration of substances which are retained in the sperm cytoplasm. it was suggested by Werner (1 966) that these nuclear derivatives may give rise to the centriole adjunct and similar structures. However, this finding calls for further verification. Stable microtubules may also be endonuclear and protrude outwards in evaginations of the nuclear and plasma membrane. This was pointed out in the Neuroptera (Baccetti et al., 196%). This condition is quite common in the

a

a)

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Fig. 6 . Two different forms of insect sperm nuclei. (A) The conical model of Bacillus, with an inner cavity containing polyribosome-like granules. ~ 9 0 , 0 0 0(From Baccetti et al, 1972b.) (B) The coiled arrangement found inMachi1is.x 90,000. (From Dallai, 1972.)

star-shaped nucleus of certain crustaceans (Decapoda), in which motile arms are found (Pochon-Masson, 1965, 1968a, b; Yasuzumi and Lee, 1966; Eliakova and Goriachkina, 1966;Anderson and Ellis, 1967),but is exceptional among insects. The scale insects, whose sperm is highly aberrant (see below), also seem to belong to this category.

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33 1

In the lower Arthropods, and in general among sea-dwelling invertebrates, the existence of a canal running within the nucleus, housing processes that are connected with acrosomal structures, is of common occurrence. As mentioned earlier, the sperm nucleus in insects is usually compact. However, in Bacillus, Baccetti and his collaborators (1 972b) describe a cribriform structure (Fig. 6A) full of polyribosomes positioned in an extranuclear space in the centre of the nucleus. C. CHEMICAL CHARACTERISTICS

It is well known that in the mature spermatozoon the nucleus consists essentially of the haploid DNA content. Atypical lines may lack DNA, as in apyrene lepidopteran sperms (Meves, 1903). In some instances, e.g. some carabids, polyploid sperms have been described (Bouix, 1963). Conversely, no RNA is present. In two insects, Philaenus and the house cricket, extrusion of ribonucleoprotein granules through the nuclear pores during the spermatid stage has been documented (Maillet and Gouranton, 1965; Kaye and McMaster Kaye, 1966). Along with DNA, proteins are contained in the sperm nucleus. These are mainly basic and consist of basic amino acids which replace the histones occurring in the nucleus of normal somatic cells and spermatogonias. Protamines, essentially consisting of arginine, lysine and histidine, chiefly studied in fish, are the most frequent. Other insects (Orthoptera and Drosophila) have, on the contrary, a category of histones particularly rich in arginine as a basic nuclear protein, which are synthesized in the cytoplasm during spermiogenesis (Das et al., 1964), using messenger RNA produced prior to meiosis (Bloch and Brack, 1964; Claypool and Bloch, 1967). Protamines and arginine-rich histones are essential for the compact nucleus of the mature spermatozoon. It was reported by Shoup (1967) that a particular Drosophila mutant being unable to achieve the transition from histones to arginine-rich histones possesses sterile sperms, with non-condensed nuclei. D. PHYSICAL CHARACTERISTICS

The important problem concerning the organization of material within the sperm nucleus, is still unresolved. Since X-ray observations in insects are not available, all information has been obtained from polarized light, fluorescence and electron microscopical investiga-

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tions. In other groups of animals we know that the nucleoprotein chains in the sperm nucleus are variably arranged, that is, ranging from an almost parallel array to haphazardly distributed chains. Working on the same material in insects, where the sperm nucleus is always exceedingly long and narrow, InouC and Sat0 (1 962, 1966) and MacInnes and Uretz (1968) came to opposite conclusions concerning Orthopteran spermatozoa. According t o the former workers, deoxyribonucleoproteins are organized into two superhelices of 8000 each, resulting from the coiling of a 1500 helix According to which consists in its turn of slender bundles (200 the latter workers, however, no superhelices exist and the DNA lies in arrays parallel t o the major axis of the sperm. The second interpretation has been substantially confirmed by Zirwer et al. (1970). Chevaillier (1 970), in an effort to resolve the controversy, proposes a scheme in which the chromosomes are oriented end-to-end along the length of the sperm nucleus. In this connection, Chevaillier mentions that Hughes-Schrader ( 1946) has reported a constant arrangement of the haploid chromosome set in Coccidae and quotes Taylor’s (1 964) work on Orthoptera, which by step-wise labelling with thymidine has revealed different levels of DNA in the sperm nucleus corresponding to a linear arrangement of chromosomes in the head.

a

a).

a

V. THE CENTRIOLAR REGION A. THE CENTRIOLE

In insect spermatids the normal orthogonal orientation of the two centrioles has been observed occasionally (Breland et al., 1966). More often either a single centriole is found (Friedlander and Wahrman, 1966), or two, as in biflagellate sperms (Fig. 7). However, the fine structure of this organelle has been established by only a few workers (Hoage and Kessel, 1968; Anderson and AndrC, 1968; Phillips, 1970b). They have the classical type of nine helicoidally arranged triplets (AndrC and Bernhard, 1964; Fawcett, 1966), comparable to those recently described by Ross (1968). Nevertheless, a more obscure problem today is the persistence of a centriole in the mature insect spermatozoon. As a matter of fact, no electron micrographs so far published show a classical centriole in a fully mature sperm and the presence of this organelle as late as the completion of spermiogenesis was denied by Phillips (1970b) in a

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Fig. 7. The centrioles (c) of the biflagellate sperm of Haemtopinus suis Both are surrounded by the adjunct centriole (ca). ~ 6 0 , 0 0 0(From . Baccetti el aZ., 1970a.)

recent discussion of this problem. At the same time, a centriole-like organelle was accurately described by Perotti ( 1970) in Drosophila sperm. However, in this case all the C tubules of the triplets are continuous with the accessory flagellar fibres (which, as will be reported presently, have a different origin) and the central pair of tubules is also present. These profiles could more easily be interpreted as the tip of the normal axial filament, where the accessory fibres may simulate triplets since they lie close to the peripheral doublets. At present, therefore, it may be assumed that a centriole does not occur in the mature insect sperm. B. THE CENTRIOLE ADJUNCT

This structure is a compact, basophilic sleeve, already known from light microscopy, characteristic of the spermatozoon in almost all insects. Called by various terms (“centriole adjunct”, which was

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coined by Gatenby and Tahmisian in 1959, is used most often), this organelle has been described in a variety of species in almost every order (see Cantacuzene’s survey (1 970)). It is generally of granular appearance (Fig. 7), with granules ranging between 160 and 320 a;it is fibrous in a few cases, e.g. in the house cricket and in Sciara (Kaye and McMaster-Kaye, 1966; Phillips, 1966a). Generally, it is more abundant in young spermatids (over 1 p 2 across) than in older ones where it becomes more compact (Breland et al., 1966). It is even more reduced in spermatozoa, where in some instances it may not be found (Phillips, 1970b). The origin of the centriole adjunct is still unknown. Werner (1965) favours the view of a nuclear origin starting from some material derived from a particular porous caudal portion of the nuclear membrane. Cantacuzhe (1970) showed that its synthesis was induced by the spermatid centriole. The aberrant multiplication of the centrioles is paralleled by a corresponding multiplication of the centriolar adjunct. Transitory contacts with Golgi cisternae and the endoplasmic reticulum are not sufficient to secure a supply of material by these organelles. Thus, the origin of the centriole adjunct is still uncertain. It seems to be responsible for the disappearance of the centriole around which it forms. Only recently has the nature of the centriolar adjunct been clarified by histochemical methods (Baccetti et al., 1969c; Yasuzumi et al., 1970; Gassner, 1970) and autoradiography (Baccetti et aZ., 1970a). There is little doubt that it consists of a ribonucleoprotein. This demonstration enabled Yasuzumi and his collaborators to propose a possible relationship between centriole adjunct and the chromatoid body of the spermatid. The ribonucleoprotein nature of the chromatoid body has long been claimed in many animals (Sud, 1961a, b; Daoust and Clermont, 1955) as well as in insects (Tandler and Moriber, 1965). But Eddy’s (1 970) recent demonstration that the chromatoid body is devoid of ribonucleoproteins, and that it arises not from the nucleus but rather from the interstices between the mitochondria (Fawcett et al., 1970) nullifies the main support to the derivation: nucleus-chromatoid body-centriole adjunct. The function of the centriole adjunct is not clear. Beginning with Gatenby and Tahmisian (1959) and Gatenby (1 96 l), most workers have always favoured a mechanical function for this compact collar, in the sense of it fastening the sperm head and tail together. This view is shared by Breland et al. (1966) and Fawcett and Phillips (1 969). Only after the more recent chemical analyses have other

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hypotheses been advanced, regarding it as a deposit of basic histones released by the nucleus during its enrichment with arginine-rich histones (Cantacuzkne, 1970) or as a concentration point of substances to be utilized in the assembly of the flagellar filaments (Yasuzumi et al., 1970). Since the initial segment of the axial filament is housed inside the centriole adjunct which apparently functions to initiate the movement of the flagellum (Baccetti et al., 1972b), a mechanical action of the centriole adjunct appears more and more probable. C. THE INITIAL SEGMENT OF THE AXONEME

Granted the absence of the centriole in insect spermatozoa, it follows that the first tubular units one encounters directly behind the sperm head are the fibres of the axial flagellar complex. In some instances, the accessory tubules are the first t o appear, in others the doublets; in all cases, the two central units arise shortly thereafter. This initial segment of the axoneme fits into ‘a corresponding nuclear indentation, or inside the sleeve formed by the condensed centriole adjunct. Its structure is complicated by the fact that it contains some units which are no longer found in the following tracts. The most widespread arrangement (Fig. 8C) consists of two short concentrically orientated cylinders connected by nine longitudinal radial laminae dividing the interspace into sectors (Fig. 4). Each doublet and each accessory tubule are contained in one sector, lying against a

Fig. 8. The “initiation motor” region of insect sperms. (A) and (B) Two different levels inDrosophiZa, showing cross links (arrows)between the opposite doublets. x 120,000. (From Perotti, 1970.) (C) Cross section in Euccillus, showing the outer ( 0 ) and the inner (i) cylinder and the radial laminae (I). ~90, 000.(From Baccetti et aZ., 1972b.)

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lamina. The two central units are contained within the central cylinder. Tail sections in different functional moments show that laminae and cylinders lie at variable angles and the laminae themselves differ in their extension. These laminae are strongly reactive for ATPase activity. Baccetti and his collaborators ( 1972b) have recently proposed that this complex may function as a device to trigger the rotation of the inner cylinder within the outer one, thereby dragging the heads of the axoneme tubules into this motion, so as to initiate a wave which then propagates along the fibres (Section X A). An arrangement of this type is evident morphologically in Phasmoidea, Orthoptera, Dermaptera and Neuroptera. In the spermatozoa of Drosophilu, the two cylinders are not present in the neck region (Perotti, 1970), but cross links are evident between the opposed triplets (doublets plus accessory fibres) each embedded in a dense sheath (Fig. 8A, B). In the coleopteran Tenebrio, both the inner cylinder and the radial or cross laminae (or strands) are missing, while a thick outer cylinder is found, which connects the heads of the outer fibre crown one after the other. In point of fact, the wave pattern in the three arrangements is considerably different (see Section X A). The organization of the axoneme is undoubtedly very complex and is relevant to the general problem of flagellar motion. In a schematic diagram of a normal cilium at different levels, Gibbons and Grimstone (1 960) and Holwill(l966) illustrate a structure composed of two cylinders and radial laminae in proximity to the head, followed by another consisting of a single outer cylinder and one more with cross strands. As a working hypothesis, it can be assumed that from the primitive multiphase mechanism occurring in cilia one stage or another was preferentially developed by spermatozoa, resulting in the formation of different kinds of waves. VI. THE AXIAL FLAGELLAR FILAMENT OR AXONEME

By slightly modifying Warner’s definition (1970), the axoneme (Fig. 9) may be regarded as the whole of the flagella complex limited by the outermost crown of microtubules. Hence, the classical Fig. 9(A) Cross section of sperm tails of Ceratitis capitata (Diptera): a,axoneme; m, mitochondrial derivatives; p, plasma membrane. ~120,000.(B) Cross section of the

sperm tail of Tenebrio molitor (Coleoptera): ab, accessory bodies; at, accessory tubules; cf, coarse fibres; ct, central tubules; d, doublets; Ih,link heads; md, mitochondrial derivatives. ~240,000.(From Baccetti et aZ., 1972a.)

Fig. 9

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microtubules, arranged according to the fundamental 9 + 2 or 9 + 9 + 2 patterns, along with the elements connecting them with one another contribute to its formation, while both mitochondria and membranes are excluded. A. THE MICROTUBULES

In the vast majority of insects, the axoneme consists of two central tubules, nine doublets and nine accessory tubules (Figs 4,9B, 10A, D, E). This scheme was first brought to light by Rothschild (1955) in the honey-bee and by Yasuzumi (1956) in Drosophila. It was then established that most insect orders belong to this category. The only exceptions are the Protura (Fig. 12A), which have a bizarre axoneme with 12 (Acerentulus) or 14 (Acerentomon) doublets and nothing else (Baccetti et al., 1972d), and the other primitive Apterygota (Collembola and Japigidae), in which the accessory tubules are missing and the basic 9 + 2 pattern, typical of classical primitive sperms, is retained (Krzystofowicz and Byczkowska-Smyk, 1966; Dallai, 1967, 1970; Baccetti and Dallai, 1972). Accessory tubules are also lacking (Fig. 10B) in the two kindred orders Mecoptera (Baccetti et al., 1969c) and Aphaniptera (Baccetti, 1968; Phillips, 1969), as well as in two species of Trichoptera (Phillips, 1970b) and in Thysanoptera (Baccetti et al., 1969d). Other arrangements have occasionally been reported, e.g. the Ephemeroptera (Fig. 1OC) lack both central tubules, thus being 9 + 9 + 0 (Baccetti et al., 1969b; Phillips, 1969); the same is true of a Psocid, which however has a solid central fibre (Phillips, 1969). Culicid Diptera, on the contrary, have only one central tubule and are thus 9 + 9 + 1 (Breland et al., 1966). A non-identified mycetophilid dipteran has three central tubules, hence is 9 + 9 + 3 (Phillips, 1970b). The two trichopteran species mentioned above have as many as seven central tubules being therefore 9 + 7 (Phillips, 1969). The dipteran Sciara coprophila (Fig. 11) lacks central tubules, having instead 70 doublets and 70 spirally arranged accessory tubules in its huge flagellum (Makielski, 1966; Phillips, 1966b). In general, therefore, there are three kinds of tubules in the axoneme of insect spermatozoa: accessory, doublets and central tubules. These are most often present in the 9 + 9 + 2 pattern. Differences among the various tubule categories have already emerged from structural studies (Andre, 1961; Baccetti and Bairati, 1964) and were conclusively demonstrated by Behnke and Forer

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Fig. 10. Cross section of five different sperm tails: ab, accessory bodies; at, accessory tubules; ct, central tubules; d, doublets; md, mitochondria1 derivatives. (A) Neuronia 9 + 9 + 2 (Trichoptera). ~130,000. (From Phillips, 1970.) (B) Panorpa annexa, 9 + 2 (Mecoptera) ~ 9 0 , 0 0 0 .(From Baccetti et al., 1969c.) (C) Chloeon dipteron, 9 + 9 + 0 (Ephemeroptera). ~120,000.(From Baccetti et al., 1969b.) (D)Noronectuglauca, 9 + 9 + 2 (Rhynchota). ~90,000. (E) Chrysopa curnea 9 + 9 + 2 (Neuroptera). ~90,000.(From Baccetti e t al., 1969c.)

Fig. 11. Cross sections of the sperm tail of Sciuru coprophila. ~45,000.(From Phillips, 1966b.)

( 1967) by means of differential enzymatic extractions upon ultrathin

sections. The following knowledge is now available about this subject.

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34 1

1. Central Tubules a. Tubules. The diameter of central tubules from different axonemes

varies from 200 to 300 A. Their wall is about 70-80 A and its globular structure was first demonstrated by Bairati and Baccetti (1965). After dissociation it resolves into subunits appearing as fibrils made up of an alignment of globules or microcylinders, some 40 A thick. As a rule, the wall of each tubule consists of about 10 subunits. In various mammals, AndrC and ThiCry (1963) and Pease (1963) counted 10 of them, but among insects their number seems a little higher, i.e. 13 (Fig. 9B), according to the measurements of Phillips (1966), which were carried out on many species from different orders. There are 10-12 according to Danilova (1 969) in Bombyx mori. It is important to recall that 12-13 subunits are found in cytoplasmic microtubules and in those of the flagellum of protozoa (Ledbetter and Porter, 1964; Gall, 1965, 1966; Behnke and Zelander, 1967; Ringo, 1967; Fuge, 1968). In insects, no detailed chemical investigation has been performed'on the tubule wall. It is,

however, known to be proteinaceous (Behnke and Forer, 1967); possibly the tubulin of Mohri (1968). The latter is an actin-like protein with a 6 s sedimentation constant and molecular weight of about 120,000 changing to about 60,000 after denaturation. It contains a guanine nucleotide and binding sites for colchicine (Shelanski and Taylor, 1968). The above protein was studied in echinoderm sperm (Plowman and Nelson, 1962; Nelson, 1966; Shelanski and Taylor, 1968) as well as in the doublets of Tetrahymena cilia (Stevens et al., 1967; Renaud et aZ., 1968). The biochemical unit (mol. wt = 120,000) is likely to correspond to two of the 40 a microcylinders described by morphologists (Shelanski and Taylor, 1968). According to Fine (1971) it seems to be composed of two different subunits, with mol. wt. = 56,000 and 54,500 respectively, with different amino acid composition (Bryan and Wilson, 1971). These subunits would then be able to aggregate into longitudinal fibrils or to follow a helical course. This explains the different appearance shown by the same microtubules when examined longitudinally after dissociation in negative stain (Danilova, 19691, while in cross section they always look the same (Thomas, 1970; Henley, 1970). The central tubules may be hollow (Fig. 10B, E), as in Mecoptera, Plecoptera and Neuroptera (Baccetti et aZ., 1969c, 1970e) or may contain a globular central core (Fig. 9), of the same size as the wall subunits, as is the case among Diptera (Bairati and Baccetti, 1965; Phillips, 1966b; Perotti, 1969; Warner, 1970). In some cases, they may be filled with microcylinders, some

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Fig. 12(A) Cross section of the 14 + 0 axoneme of Acerentomon majus (Protura). ~80,000. (From Baccetti et al., 1972d.) (B) Cross section of the 9 + 9 + 2 sperms of Campodeu (Diplura) showing the disordered migration of accessory tubules near the mitochondrion (arrows). ~48,000.(From Baccetti and Dallai, 1972.) (C)A more caudal section of the 9 + 9 + 2 sperm of Campodea (Diplura) showing the ordered disposition of the nine accessory tubules (arrows) near the mitochondrion. ~75,000.(From Baccetti and Dallai, 1972.) (D) The tip of the sperm tail in Telamona spec. (Rhynchota). The doublets appear dissociated. ~62,000.(From Phillips, 1970a.)

40 A in diameter, about eight per tubule, as in Gryllus (Kaye, 1964, 1970). Very often (in Trichoptera, Lepidoptera, Phasmoidea), however, the two tubules (Fig. 14A, B, D) become filled not only by protein, but also by a glycogen-like polysaccharide (Baccetti et a l , 1969a, 1970e, 1972b). b. The projections. Short side-projections (Kessel, 1967) are seen to jut out from the tubule wall, 150-170 A apart from one another and helically coiled (Fig. 13B, C) around the tubules (Perotti, 1969). In the cilia and flagella of Protozoa these projections exist in two series in tubule I and in one series in tubule I1 (Hopkins, 1970), or are present only in tubule I (Chasey, 1969). They seem to be separated from the tubule wall and vertically interconnected by filaments (Chasey, 1969).

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According to Warner (1970) they might be the binding site of the tubule wall to the “central sheath” which is also helically coiled around both structures with a similar pitch of 160 A . The central tubules do not generally undergo transformations towards the tail end, where they usually terminate after the accessory tubules and shortly before the doublets. In a few instances, e.g. Drosophila and the Trichoptera (Perotti, 1969; Baccetti et al., 1970e) they end after the doublets. Rarely, as in Tenebrio, they are the first to disappear (Baccetti et al., 1972a). In Gryllotalpa they are filled with a glycoprotein which becomes tanned, thereby making the distal half of the tail rigid and stiff (Baccetti et al., 197 1b). 2. Doublet tubules

a. The tubules. These are the only structures which are consistently present in all types of flagellate spermatozoa. Doublets are, as a rule, nine in number, lying at some distance from one another. They can be oriented and numbered from 1 to 9 .according to the general pattern of cilia and flagella (Afzelius, 1959; Gibbons and Grimstone, 1960): so that doublet 1 lies on the median plane normal to the line linking the two central tubules. Doublet 2 follows in the arm direction and so on. Only Protura exhibit 12 or 14 doublets (Baccetti et al., 1972d); in Sciara there are 70 (Phillips, 1966a: Fig. 11). In cross section both tubules in each doublet (Fig. 9B) are found to consist of a B tubule (200-230 A ) slightly larger than the A tubule (200 A). From tubule A two arms are seen to extend. The outer one is about 35 mp long and directed towards the flagellum centre, the inner one is only 20 mp long. These measurements have been checked by Allen (1968) in Tetrahymena cilia and seem to be valid for the insect flagellum (Warner, 1970). The arm pairs, up to 100 in thickness, are found every 200-220 A along the tubule length (Phillips, 1970b). The doublet wall and the arms are not made up of the same material. Both are proteinaceous (Fig. 14C) but, like the central tubules, the doublet consists essentially of tubulin (although this evidence concerns protozoan cilia and sperms from Echinodermata as emerges from studies carried out primarily by Renaud et al. in 1968 and by Shelanski and Taylor in the same year). The arms, however, consist of dynein, a 14s protein unit with a molecular weight of 600,000, arranged in linear polymers endowed with ATPase activity (Fig. 14E). These observations are based on studies of Tetrahymena and the sea urchin by Gibbons (1963) and Gibbons and Rowe (1 965). It is most unlikely that the sperm doublets differ

a

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Fig. 13

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345

from this structure. Unlike the tubule walls, the arms of the A tubules are rich in ATPase activity (Baccetti et al., 1972b). Both consist of globular units of comparable size with those of tubulin. However, they differ from one another in features other than their B is not a real size. Firstly, as shown by Phillips ( 1 9 6 6 ~ )tubule ~ tubule, but a groove adhering to tubule A. Nevertheless, the subunit number seems to be consistently 13 and it may be supposed that B opens against A. Furthermore, a chemical difference between the two tubules has emerged from enzymatic extractions of various types as reported by Behnke and Forer (1967), though in both cases they are proteins (Baccetti et al., 1970e). As a rule, tubule A appears “solid” and tubule B “hollow”. When, as in Gryllus, both are filled up with globular units, these are in greater number in A than in B, though the latter is narrower (Kaye, 1970). In Bacillus spermatids tubule B has glucose-6-phosphatase activity (Baccetti et al., 1972b). A new technique of heat fractionation devised by Stephens (1970) separated the doublets of sea urchin spermatozoa into two units. He was thereby able to isolate A and B tubules which had a similar guanine nucleotide content, but different levels of amino acids, in particular cystein. This explains why only tubule A can bind to dynein and raises some questions about other microtubule categories. The contractility problem also remains open since, again in the sea urchin, tubulin does not possess antigenic properties like those of actin (Stephens, 1970). Recently, Behnke et al. (1971) were able to demonstrate in Nephrotoma sperm tails, bundles of 15-20 actin filaments independent of the axoneme tubules. This actin may be involved in sperm motility, but its exact localization in the sperm is still unknown. The doublets are usually the longest sperm tubules and are the last to terminate at the tail tip. Often they dissociate into their two units (Phillips, 1 9 6 6 ~ ) In . the Homoptera the tail splits into four strands (Fig. 12D), each containing some doublets, which eventually dissociate (Folliot, 1970). In Gryllotalpa the doublets are invaded in Fig. 13. Fine structural pattern of the insect axoneme. (A) Radial links (RL) and peripheral doublet (D) of Drosophila, negatively stained. The doublet (D) appears not dissociated. ~ 140,000.(From Bairati and Perotti, 1970.) (B) Central tubule of Drosophila, negatively stained. Projections spaced at 170 A are evident. ~200,000.(From Bairati and Perotti, 1970.) (C) Platinum shadowed central tubule of Aiolopus. A helicoidal surface pattern is evident. ~ 48, 000.(D) Acid phosphatase in Ceratitis (Diptera) spermatozoa. The activity is evident on the plasma membrane and in the matrix surrounding the tubules. ~120,000.(E) Glycogen on the link-heads in cumpodea (Diplura). Thi&y’s method. ~75,000.(From Baccetti and Dallai, 1972.)

346

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Fig. 14

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the distal half by a glycoprotein material which undergoes tanning, thus stiffening the entire tail segment (Baccetti et al., 1971b). In a few cases (e.g. in Trichoptera and Drosophila), the tubules are the first to end among the axonemal tubules (Baccetti et al., 1970e; Perotti, 1969). b. The radial links. Besides arms, the doublets also possess other projections which are directed centripetally (Fig. 4). Cross-sectional profiles suggested that they were radial laminae, which was, in fact, the earliest interpretation. They are radial links consisting of straight rays some 325 A long and 70-125 A thick (Behnke and Forer, 1967; Perotti, 1969; Warner, 1970) more or less orthogonally oriented to the fibre and alternatively spaced at 320 A and 560 A . When viewed longitudinally, each closely adhering pair is separated by a wide interval from the other. On the two opposite sides of the flagellum, the radial links are staggered 200 A from each other in a helical coil (Warner, 1970). An exactly comparable situation was reported in Chlamydomonas (Hopkins, 1970), hence it may possibly be of general occurrence.

3. Accessory Tubules These are characteristic, apart from a few exceptions, of the most evolved insects, from Campodeidae Diplura upwards. Their structure and size are closely reminiscent of those of the two central tubules (Fig. 9A, B). However, in some species they are thinner, while in other species (Trichoptera, Lepidoptera) they are thicker (over 300 A in diameter). In cross-section they show 13 globular units in their wall as in the two central tubules (Phillips, 1 9 6 6 ~ )Where . their diameter is wider, these units are in greater numbers; for instance in a Fig. 14. Histochemistry of the tail of insect spermatozoa. (A) Glycogen (arrows) in the central and accessory tubules of Bacillus rossius sperm. ThiCry’s method. ~75,000.(From Bigliardi et Q L , 1970.) (B) Glycogen (arrows) in central and accessory tubule ofhlystacides azurea (Trichoptera) sperm. Thikry’s method. ~90,000.(From Baccetti et al., 1970e.) (C) Pepsin treated sperm tail of Nemoura cinerea (Plecoptera). Doublets and mitochondria1 derivatives (arrows) appear significantly extracted. ~30,000. (From Baccetti et at., 1970e.) (D) Amylase treated sperm tail of Ceratitis capitata (Diptera). Only the accessory tubules (From Bigliardi et al., 1970:) and the two central ones (arrows) appear extracted. ~60,000. (E) ATPase in the sperm tail of Bacillus rossius (Phasmoidea). The reaction (arrows) 1s positive on the coarse fibres, on the arms of the doublets, on the central sheath and on the accessory bodies. ~60,000.(From Bigliardi e t al., 1970.) (F)UTPase in the same material. The reaction appears similar to that of ATPase. ~60,000.(From Bigliardi et al., 1970.) (G) UTPase in the sperm tail of Ceratitis capitata (Diptera). The reaction is positive only in axonema1 (coarse fibres, arms, central sheath) structures. ~60,000.(From Bigliardi et a/., 1970.)

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Coleopteran, as many as 15 or 16 were counted by Shay et al. (1 969). During spermiogenesis, these tubules arise from laminar outgrowths of the B tubules of the doublets (Cameron, 1965). These projections (Fig. 25A, B) first extend outwards in the form of a groove, then bend and once the tubular shape is achieved they separate from the doublets. Later, the material filling them makes its appearance. Therefore, their major chemical component should resemble that of the B tubules, i.e. tubulin B, but no information about this is available. When viewed longitudinally, projections are also discernible which may possibly connect adjacent accessory tubules. The content of the accessory tubules often resembles that of the central ones. In some species which have hollow accessory tubules, the central ones are hollow as well, e.g. in Plecoptera (Baccetti et al., 1970e). In other species (e.g. among the Diptera mentioned earlier) a globular osmiophilic core is found in both categories (Fig. 9). In rare cases, the central tubules are hollow (Fig. lOE), while an osmiophilic core is found in the accessory ones (Neuroptera: Baccetti et al., 1969c). The invasion by microcylinders, typical of Orthoptera (Kaye, 1964, 1970), is more massive in the accessory tubules; up to 15 units can be found in each tubule. A similar event was also reported in Coleoptera (Cameron, 1965; Baccetti, et al., 1972a) where there are fewer, namely, 6-7 units. The most spectacular behaviour is provided by the accessory tubules containing the glycogen-like polysaccharide (Phasmoidea, Trichoptera, Lepidoptera) as reported by Baccetti et al. (1969a, 1970e, 1972b). This polysaccharide appears as a compact mass which fills up the huge tubules previously described as over 300 A in diameter (AndrB, 1961; Yasuzumi and Oura, 1964). In the spermatid stage, during which the accessory tubules are being filled, nine Golgi vesicles (Fig. 25A, D) surround the axoneme, each concentrating around one tubule. Transfer of material, however, could not be established (Baccetti et al., 1972b). In the flagellum of some species, these accessory tubules arise more apically than the other axonemal units (Phillips, 1 9 6 6 ~ ) Since . they are secreted by the B tubules, they are pushed forward after separating from the latter. As a rule, these accessory tubules terminate towards the tail tip before the doublets and the central units. In a few cases, however, such as Trichoptera or Tenebrio (Baccetti et al., 1970e, 1972a), they are the last ones to end.

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In Diplura Campodeidae a 9 + 9 + 2 arrangement is differentiated early in the spermatid. The accessory tubules soon migrate around it and are arranged in two crescents (Fig. 12B, C), one consisting of five and the other of four units aligned against doublets 1, 7 and 8 on one side only of the axoneme. They are separated by the mitochondrion which is juxtaposed to doublet 9 (Baccetti and Dallai, 1972). Similar behaviour is shown in a thysanuran (Machilidae) in which the two crescent-shaped accessory bodies which flank the axoneme displace the accessory tubules into two series, one consisting of four and the other of five elements (Dallai, 1972). The classical 9 + 9 + 2 arrangement is first found in the highest Thysanura, namely, the Lepismatidae (Bawa, 1964; Werner, 1964), and is retained thereafter by all the Pterygota. B. THE CENTRAL SHEATH

This is a structure which surrounds the two central tubules (Fig. 4). In cross-sectional view it shows a circular profile about 700 A in diameter and ranges in thickness from 150 A , in the sector facing doublet 1, t o 90 4 in the contralateral one facing doublets 5 and 6 (Warner, 1970). In longitudinal sections it appears as a helically coiled band, with a pitch of 120 A (Andre, 1961) or of 160 A (Warner, 1970). This pitch, equal to the period of the projections of the central tubules, suggests that the central sheath is somehow anchored to it, even though the latter does not seem to run, as suggested by longitudinal sections, in exact correspondence to the lateral edges of the central tubules. The central sheath is ill-defined from a chemical standpoint because it is an extremely labile structure difficult to isolate or examine by negative staining. How it comes to be connected to the heads of the radial links remains obscure. It certainly contains (Fig. 14E, F, G) ATPase and UTPase (Baccetti et al., 1972b) and is present when, as in Ephemeroptera (Fig. 1OC), the central tubules are completely absent (Baccetti et al., 1969b). C . THE LINK-HEADS

The presence of nine fibrous units between the central tubules and the doublets was demonstrated in the Flagellates by Gibbons and Grimstone (1 960), who called them secondary fibres. Afzelius (1959), in the sea urchin sperm, interpreted them as juxtaposed

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radial links. But their nature remains obscure. Other elements in favour of the existence of longitudinal filamentous connections between the thickenings of the links lying in the same row, have been provided by Andre (1 96 1) in insect spermatozoa, and by Birge and Doolin (1 969) in vertebrate cilia. Almost all other workers have disregarded this problem. More recently, the presence of secondary longitudinal fibres has been disproved both in the flagellate flagellum (Hopkins, 1970) and insect sperms (Perotti, 1969; Bairati and Perotti, 1970; Warner, 1970) as well as in mammalian and amphibian sperms (Fawcett, 1970). All the data available in the literature related to the secondary fibrils in spermatozoa must be ascribed to the link-heads. The link-heads were first described by Andr6 (1961). In cross section, it was shown by Bairati and Baccetti (1965) that they are bipartite; this finding has been consistently confirmed by careful measurements done recently by Warner (1 970): in cross section they measure 200 A by 280 A . This value (Fig. 9A, B) is valid for the Diptera, a Dipluran Cizmpodea, Coleoptera and Orthoptera (Baccetti and Dallai, 1972; Baccetti et al., 1972a). However, a second arrangement exists (Fig. 24A) in Phasmoidea (Baccetti et al., 1972b) and in another Dipluran, Japyx (Baccetti and Dallai, 1972) in which the bipartite structure is less evident and the link-head is smaller in diameter, i.e. about 150 A by 180 A . They are chemically and structurally rather labile, hence poorly understood. Baccetti and Dallai (1 972) showed histochemically the presence of glycogen (Fig. 13E) at the same sites as both the large and small forms of link-heads in Diplura, while such glycogen deposits are not found in all other insects (see Anderson and Personne, 1970). A similar distribution of glycogen was described in some Gastropoda (Anderson and Personne, 1970). D. THE COARSE FIBRES

In the typical insect spermatozoon, an important material exists between the accessory tubules, which is rarely studied because it is difficult to fix and preserve. Baccetti ( 1963) and Baccetti and Bairati (1964) were able to identify four components in what were called “outer fibres” (Andr6, 1961) in Drosophila and Dacus, one of which obviously corresponds to the accessory tubule. Later, Bairati and Baccetti (1965) after identifying the accessory tubules, regarded the interposed material as amorphous, but viewed the presence of the interposed “X granule” as a unit in itself. Generally, all the material

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lying between the accessory tubules was considered as amorphous by later workers and neglected by them. However, Daems et al. (1 963), as well as Bigliardi et al. (1970), found ATPase and also UTPase localized in it (Fig. 14E, F, G). In Sarcophaga, Warner (1970) has recently identified two coarse fibres lying between the accessory tubules, thus confirming an interpretation similar to that of Baccetti and Bairati (1964). On the basis of histochemical data partly obtained from Dipteran spermatozoa, both coarse fibres (Fig. 14E, F, G) had ATPase and UTPase activity. However, they are certainly not the same in all the insects. On the basis of the literature on this subject it emerges that in Diptera (Fig. 9A) the coarse fibres are generally as Warner (1970) described them and the same is true for Coleoptera (Baccetti et al., 1972a: Fig. 9B) and Phasmoidea (Baccetti et al., 1972b) (Fig. 24A) and even in Thysanura Lepismatidae (Bawa, 1964). In other cases (Fig. lOE), however, they are much attenuated (Neuroptera, Lepidoptera, Plecoptera) and occasionally (Ephemeroptera: Baccetti et ul., 1969b) entirely absent (Fig. 1OC). A peculiar case is found in Diplura Campodeidae in which

Fig. 15. Biflagellate spermatozoa of Rhynchotoidea. (From Baccetti et aZ., 1969d.) (A) The sperm tail ofMenopongallime (Mallophaga).Two axonemes (a) and two mitochondrial (m) derivatives are evident. ~ 6 0 , 0 0 0 .(B) The sperm tail of Chryptothrips latus (Thysanoptera). The axoneme is composed by 18 doublets and four single tubules.

~135.000.

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the accessory tubules are all assembled against doublets 1, 7 and 8 without intervening material. External to doublets 2, 3, 4, 5 and 6, however, there are a series of associated osmiophilic structures which look like coarse fibres and are not separated by tubules. In the case of Bacillus spermatozoa, various nucleoside-phosphate phosphatases (GTPase, ADPase, CTPase) were identified biochemically in the same fraction (Bigliardi et al., 1970). It may be speculated that these enzymes are also concentrated in the same zone together with ATP- and UTPase, thus being involved in flagellar motility . E. THE AXONEMAL MATRIX

This is represented by all the space lying between the structures listed above and separated from the compartment in which the mitochondria1 derivatives are located, by means of membrane residues originating from the Golgi. This space often appears empty in electron micrographs. However, it contains a number of enzymatic activities detectable histochemically or by means of biochemical assays of the supernatant fraction. Histochemically, the most prominent enzyme (Fig. 13D) in this space is acid phosphatase (Bigliardi e t al., 1970; Baccetti et al., 1970b). Lactic dehydrogenase and phosphorylase are also abundant. They may also be detected on the plasma membrane, though only on the inner cytoplasmic membranes (partly plasma membrane, partly Golgi-derived ones) surrounding the axoneme. These enzymes are concerned with carbohydrate metabolism. Biochemical assays carried out on lactic dehydrogenase show that it is essentially present in the supernatant fraction from an Orthopteran, Aiolopus (Baccetti et al., 1970c) while it is strongly associated with membranes in a Phasmoid, Bacillus rossius, where it is quantitatively more important (Baccetti et al., 1972b). The different localization in the two foregoing instances may indicate a different physiological role, bearing in mind that Aiolopus sperm is capable of aerobic metabolism, while that of Bacillus is devoid of mitochondria, hence of cytochrome oxidase. Other enzymes involved in carbohydrate metabolism, i.e. phosphofructokinase, fructose-diphosphate aldolase and glyceraldehyde-3phosphate-dehydrogenase were identified in Bacillus rossius spermatozoa (Baccetti et al., 1972b). So far, their localization is unknown. This enzyme assembly, along with the presence of a polysaccharide in the accessory tubules, points to the existence of a glycolytic

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Fig. 16. Cross sections of several spermatozoa of the Coccid insect Parlatoria aleae: c, chromatin; m, microtubules; n, nucleus. ~70,000. (From Robison, 1970.)

pathway in the axonemal compartment. As a rule, carbohydrate utilization goes t o completion in the reactions catalyzed by mitochondria1 structures. In the absence of the latter (see next Section) it may be deduced that lactic acid is the end metabolite (Baccetti et al., 1972b). AIP--15

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VII. MITOCHONDRIA

As mentioned in the introduction, one of the essential features of the insect sperm is the peculiar evolution of the mitochondria, which are transformed into elongated structures flanking the axoneme. Their organization, however, is not the same in all insects, neither is it simple nor fully understood. A few very primitive insects exist in which the mature sperm has normal mitochondria which are fused into elongated masses. In others, some impressive mitochondrial transformations have occurred, while others lack mitochondria altogether . A. NORMAL MITOCHONDRIA

We may consider as normal mitochondria those with a moderately opaque matrix free of crystalline materials containing abundant well-developed inner cristae with clear-cut cytochrome oxidase activity. This mitochondrial type (Fig. 21 E) is found only among the most primitive insect groups, namely, in all of the Collembola (Dallai, 1967, 1970) and Diplura (Baccetti and Dallai, 1972), in the primitive Machilidae among Thysanura (Dallai, 1972) and in the most primitive Pterygota (Fig. 1OC), the Ephemeroptera (Baccetti et aL, 1969b). In all these cases the mitochondria, though retaining their normal aspect, undergo complex rearrangements during the spermatid phase. At this stage, mitochondria are numerous, small in size and have a tendency to fuse into a few units, undergoing the same rearrangements that were studied in greater detail in the more evolved mitochondria. These transformations lead to three elongated mitochondria in the sperm tail of Collembola, t o two in that of Diplura and Machilidae and to one in the Ephemeroptera sperm tail. These mitochondria retain both normal structure and function throughout the life of the sperm. In some, among the most involuted spermatozoa (Fig. 27) (e.g. those from the higher Termites, Reticulitermes), the mitochondria are also of the conventional model, namely small in size, numerous and provided with cristae (Baccetti eta]., 1972e). B. MITOCHONDRIA TRANSFORMED INTO DERIVATIVES WITH A CRYSTALLINE CORE

This, the best known mitochondrial category, was first studied by light microscopy (Bowen, 1920). The fine structural details have

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been fully described by Andre (1959, 1962) in Lepidoptera, by Pratt (1968) in a Hemipteran as well as by occasional reports from several other authors. All this data was coordinated by Phillips (1970b) who provided a scheme which can be applied to almost all insects. During the last spermatogonial generations and spermatocyte I prophase, the small spherical mitochondria increase considerably in number, until they fuse into long chains, oriented by the poles, forming a “palisade around the spindle” (Andre, 1962). Thus, meiosis leads to an exact distribution of mitochondria between the two daughter cells, until, during the second telophase, the mitochondria pack together into a skein (Retzius’ Nebenkern, 1904). Then there begins in each spermatid, the slow and conspicuous metamorphosis (Fig. 17) of the mitochondria into the Nebenkern through stages which And& called “twinned”, “chromophobic envelope”, “onion” and “loaf”. During these rearrangements two extremely long, labyrinthically interwoven identical chondrioconts are formed (Pratt, 1968) which eventually separate and come to lie on either side of the incipient axoneme. Together with the latter, the mitochondria develop within a narrow evagination of the plasma membrane against which the nucleus is first located. A cytoplasmic space gradually (Figs 18, 19, and 20) forms between the nucleus and the periaxonemal membrane as the latter is dragged distalwards away from the nucleus by the ring centriole (Fawcett and Phillips, 1970). The two Nebenkern derivatives insert themselves within this space (Phillips, 1970b), progressively elongating and flanking the axoneme. They taper until they become equal in size to the axoneme itself. In the Pterygota there are usually two mitochondrial derivatives (Figs4, 9, and 10) which either remain alike or one of them overtakes the other in length or in width (see Phillips (1970b) for a detailed case history). Occasionally, one of them progressively diminishes until it vanishes, thus the mature sperm possesses but one mitochondrial derivative (Fig. 10A, B). This occurs in the Mecoptera (Baccetti et aZ., 1969c) and Trichoptera (Baccetti et aZ., 1970e; Phillips, 1970b). Likewise in Thysanoptera (Baccetti et al., 1969d) (Fig. 15B) and in Diptera (e.g. Psychodidae, Baccetti et al., 1972c) (Fig. 26) only one mitochondrial derivative was found. In these cases, however, the spermatid mitochondria fuse directly into a single mass rather than two initial ones. When the two mitochondrial derivatives have flanked the axoneme for almost its entire length (instances of their equalling the axoneme in length are extremely rare) (Folliot, 1970), two important changes

Fig. 1 7

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take place. First of all (Fig. 22C), the cristae become aligned, regularly spaced at right-angles to one side of the major axis of the derivative, looking like a series of short lamellae. Then within the mitochondrial matrix some masses of material contained in regular granules, with a paracrystalline structure (Fig. 21 D), make their appearance. These were studied first by Andr6 (1 962). Subsequently they were resolved by Meyer ( 1964) and by Phillips (1 966a) using negative stains. This paracrystalline material invades the space left free of cristae in all the insects so far studied. It consists (Fig. 14C) of a protein (Baccetti et al., 1969c, 1970e) whose three-dimensional organization was suspected of varying according t o species (Meyer, 1964). It contained various types of longitudinal and transverse periodicities (the most common being 450 a) and different unit arrangements, which generally are hexagonally spaced. Two diverse crystalline organizations within the same mitochondrion have been found. The example quoted by Phillips (1970b) is not relevant, however, since the mitochondria described ‘are in reality an ordered extramitochondrial material (Mazzini, 1970). Intramitochondrial crystals probably have a pattern which is very common to many, if not all, insect orders. In Fig. 22(A, B), the same structure is demonstrated in a Dipteran and in a Coleopteran sperm. This material does not arise spontaneously within the mitochondria (Baccetti et ul., 1972a). During the last phases of spermiogenesis it was noticed by Baccetti and his collaborators (1 972a) that the Golgi cisternae come into contact with the mitochondrial wall and deposition of the early proteinaceous aggregates takes place at this site (Fig. 21A, B, C). This material is of unknown significance. In several species, repeated histochemical investigations (Bigliardi et ul., 1970; Baccetti et al., 1970b, 1972a) have shown that cytochrome oxidase is confined to the short cristae (Fig. 22C) and that the pro teinaceous matrix contains none of the enzymes concerned with respiration, glycolysis or the hydrolysis of triphosphate nucleosides. In all the spermatozoa provided with mitochondria, therefore, there are two pathways for energy production in the flagellum.

Fig. 17. Various stages of mitochondrial condensation and Nebenkern formation in the spermatid of Murgantia hisrrionicu (Rhynchota). (From Pratt, 1970.) (A) and (B). Early cluster or “twinned” Nebenkern. ~18,000.(C) Later cluster or “twinned” Nebenkern. ~16,000.(D) “Small sheet” or “chromophobic envelope” Nebenkern. x 17,000. (E) “Four layered” or “onion” Nebenkern. x 16,000. (F) “Two layered” or “loaf” Nebenkern. x 17,000.

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Fig. 18. Young spermatid of Tenebrio (Coleoptera). The ring centriole (rc) is still very close to the centriole (c) whereas the Nebenkern (n) are far away (cf., Fig. 19). ~30,000. (From Baccetti et aL, 1972a.)

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Fig. 19. Later spermatid of Euchistus (Rhynchota). The ring centriole (rc) is some distance from the centriole (c) and the two Nebenkerns (nb) are nearer than in Fig. 18. ~ 1 3 , 0 0 0(From . Phillips, 1970b.)

Normal respiration in the mitochondria1 compartment is by means of the Krebs cycle and anaerobic glycolysis which occurs in the axonemal compartment at the expense of the polysaccharide contained within the accessory tubules.

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Fig. 20. A still later stage spermatid of Euchistus (Rhynchota). The Nebenkern (ntd surrounds the axoneme between the ring centriole (rc) and centriole (c) ~13,000. (From Phillips, 1970b.) C. ABSENCE OF MITOCHONDRIA

Only one insect group provided with a typical flagellum is devoid of mitochondria. It is the order Phasmoidea (Fig. 24A), in which the genera Bacillus (Baccetti et al., 1972b) and Clitumnus have been studied. The young spermatid has a moderate complement of

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36 1

Fig. 21. Mitochondria in insect sperms. In (A), (B) and (C) a system of narrow membranes (m) originating from Golgi vesicles (g) penetrates between the axoneme (a) and mitochondrial derivatives (md) of Tenebrio spermatozoon, fusing with its limiting membranes (arrows). The inner mitochondrial crystallization (c) occurs near the region of first contact with the Golgi membranes. ~90,000.(From Baccetti et al., 1972a.) (D) Frozen-etched preparation of Tenebrio spermatozoon. The crystalline material filling the mitochondrial derivatives is evident (arrows). ~60,000. (From Baccetti et aL, 1972a.) (E) A normal mitochondrion (m) in a mature insect sperm (Campodea). ~48,000.(From Baccetti and Dallai, 1972.)

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Fig. 22. Crystallized mitochondrial derivatives in insect sperms. (A) Crystallized material of Drosophila (Diptera) after phosphotungstic acid negative staining. ~260,000.(From Bairati and Perotti, 1970.) (B) Crystallized material of Tenebrio (Coleoptera) after uranyl acetate negative staining. ~306,000.(From Baccetti et aL, 1972a.) (C) CytochromeC oxidase in the peripheral region (containing cristae) of Tenebrio mitochondrial derivatives (arrows). The crystalliie region (c) is negative. ~60,000. (From Baccetti etaZ., 1972a.)

mitochondria (which show no tendency to evolve into a Nebenkern), which possess inner cristae rich in cytochrome oxidase (Fig. 24B). At this stage, cell respiration is conducted normally. Subsequently, mitochondria undergo degeneration and are definitely absent in the spermatozoon. Biochemical assays showed that both cytochrome oxidase and succinic dehydrogenase activities were extremely low (Bigliardi et al., 1970) and were probably the result of contamination by spermatids. Oxygen consumption in sperm preparations, as

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determined in the presence of cytochrome c and ascorbate, is also absent (Baccetti e t al., 1972b). As reported in the preceding section, metabolism in the axonemal compartment is definitely reduced to glycolysis taking place under strictly anaerobic conditions. Both mitochondria or any form of mitochondrial derivative are absent from the category of non-flagellate sperms (Termites of the genus Kalotermes and coccids). In both cases (Figs 27 and 28), the spermatozoon is supplied with a rich microtubule complement (see below). In the sperm of coccids, which are the best studied, ATPase is present (Moses, 1966). Therefore, an active energy source, though different from oxidative phosphorylation, may be postulated (Ross and Robinson, 1969). In all these cases, the solution of the energetic problem may be analogous to that found in Phasmoidea spermatozoa. However, this is still an open question.

VIII. ACCESSORY ORDERED FLAGELLAR BODIES

One of the essential features of the sperm tail in insects is the fact that, besides the mitochondrial derivatives, the axoneme is flanked by one or more elongated accessory structures of paracrystalline texture. These structures are reminiscent of the compact protein core which often invades the mitochondrial matrix in Pterygotes. However, some do not seem to be related to them, either chemically or functionally. In addition, several categories may be identified.

A. STRUCTURED BODIES FLANKING NORMAL MITOCHONDRIA

This condition occurs only in the most primitive insects, in which mitochondria do not undergo their characteristic metamorphosis. In some Thysanura (Dallai, 1972) two long crystalline rodlets were described (Fig. 23B), which flank the flagellum throughout its length. They have a crescent shape when viewed in cross section. In longitudinal sections they display a paracrystalline texture with a periodicity of 200 A . In Ephemeroptera two elongated crystalline bodies (Fig. lOC) are also seen to flank the single mitochondrion. They have an average periodicity of 160 A . They seem to arise in close contact with the Golgi cisternae (Baccetti et aZ., 1969b). Their function and chemical composition is unknown.

3 64

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Fig. 23. Patterns of accessory flagellar bodies in insect sperms. (A) Tenebrio molitor (Coleoptera). ATPase present around the two bodies (b) and absent from mitochondrial derivatives (m) ~120,000.(From Baccetti et al., 1972a.) (B) Muchilis distincta (Thysmura). The accessory bodies (b) are much larger than mitochondria (m). ~ 6 0 , 0 0 0(From . DdS, 1972.) (C) Cixius netvows (Rhynchota). The accessory bodies (b) are as developed as the mitochondrial derivatives (m) ~35,000.(From Folliot and Maillet, 1970.) (D) Megouru viciue (Rhynchota). Spermatid: The accessory bodies (b) originate in contact with Golg membranes (g). ~ 6 0 , 0 0 0(From . Mazzini, 1970.)

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€3. STRUCTURED BODIES FLANKING THE MITOCHONDRIAL

DERIVATIVES WITH A CRYSTALLINE MATRIX

I. Connected with the Centriole Adjunct These formations occur in greater number and are better known than the preceding ones. In the spermatid, they make their appearance first as caudal expansions of the centriole adjunct. As they become progressively enlarged and surrounded by microtubules, they flank the axoneme and mitochondrial derivatives and end up embedded in the flagellum. A single one is found in Plecoptera (Baccetti et al., 1970e) and in Cicindela (Werner, 1965). Two symmetrical ones are present in Tenebrio (Baccetti et al., 1972a) and in Aphaniptera (Baccetti, 1968). They are primarily of a protein nature, with a carbohydrate core, demonstrated histochemically in Tenebrio. Again in Tenebrio, these structured bodies (Fig. 23A) are seen to be rich in ATPase and UTPase at least in a cortical halo consisting of globular units of 80 A . They show a compact organization in their central zone where the polysaccharide is located. The function of these formations still has to be assessed; however, the occurrence of ATPase seems to suggest some role in flagellar motility. Their origin is also unknown. The Golgi-derived cisternae never come into contact with them (Baccetti et al., 1972a). In Cicindela, Werner (1965) has documented the origin of the centriolar adjunct and its derivative from a caudal zone of the nucleus, where the nuclear membrane is highly porous. 2. Products of the Golgi Complex This section deals with the origin of the two accessory formations found in Homoptera , which are symmetrically positioned, like the mitochondrial derivatives, on either side of the axoneme. They often exhibit bizarre profiles (Fig. 23C) and a crystalline texture. They were studied in Delphacids by Herold and Munz (1967), in Cicadellidae by Phillips ( 1970b), in several Auchenorhyncha by Folliot and Maillet (1970) and in Aphids by Mazzini (1970). Their chemical characteristics and function are obscure. Only in Aphids has their origin from the Golgi-derived cisternae (Fig. 23D) been ascertained (Mazzini, 1970). C. STRUCTURED BODIES REPLACING MITOCHONDRIA

This is an extreme event typical of Phasmoidea (Bigliardi et al., 1970; Baccetti et al., 1972b). In this order, mitochondria disappear

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Fig. 24(A) The sperm tail of Bacillus rossius: ab, accessory bodies; at, accessory tubules; cf, coarse fibres; ct, central tubules; d, doublets. Mitochondria are lacking. x 120,000, (From Baccetti e t al., 1972b.) (B) Cytochrome-C-oxidasein a normal mitochondrion (m)of the spermatid of Bacillus. Seligman et al. 's method. ~ 6 0 , 0 0 0(From . Baccetti et al., 1972b.)

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during spermiogenesis and are replaced by two large bodies (Fig. 24A). These consist of alignments of proteinaceous material arranged in periodically juxtaposed small laminae and longitudinal filainents. In all sections, along almost the entire length, they appear J-shaped. They arise from the Golgi at the spermatid stage (Fig. 25B, C), are made up of proteins and are extremely rich in ATPase and UTPase. Their role is important as regards the type of locomotion, which in this case is peculiar as it follows almost circular trajectories. Although they do not appear to be structured in alternative fashions according to their thickness and do not change organization when the spermatozoon is naturally or experimentally relaxed, it was supposed that they function in contraction in view of their enzymatic equipment (Baccetti et al., 1972b). IX. SPERMATOZOA POSSESSING A DOUBLE FLAGELLAR APPARATUS OR BEING DEVOID OF IT

In relation to the rule of 9 which seems. to dominate doublet arrangement, mention has already been made of the huge flagellum of Sciara coprophila studied by Phillips (1966a) and by Makielski (1966). This has 70-90 doublets (Fig. l l ) , each flanked by an accessory tubule, which are derived from a large centriole spirally coiled around a single mitochondria1 derivative. This case seems to depend on the particular type of spermatogenesis (Metz, 1938) which leads ultimately to a single spermatid with a chromosome complement, two Xs and one or two heterochromatic chromosomes, corresponding t o the germinal tissue. In other sperm categories, described earlier, either the absence or an excess number of both central and accessory tubules can be found. A different case is represented by germ cells which either have two normal flagellar devices or where the flagellum is completely absent. A. THE PAIRED SPERMATOZOA

This is a peculiar situation found occasionally in the animal kingdom. It is generally known in Thysanwa Lepismatidae (Bawa, 1964; Werner, 1964) as well as in the coleopteran Dytiscus marginalis (Ballowitz, 1895), the gastropod Turritella terebru (Retzius, 1906; Idelman, 1960) and in American marsupials (Biggers and Creed, 1962; Phillips, 1970c, d). In Thysanura Lepismatidae, membrane fusion unites sperms two by two at the level of the nucleus.

Fig. 25

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Movement is possible only under this condition and isolated sperms are non-motile. In these insects, unlike Dytiscus, the sperm pair remains united in the female genital tract. What happens when the sperm penetrates the egg is obscure (Bawa, 1964). The type of fusion of the membranes at the nuclear level also requires further investigations. Phillips ( 1 970c, d) has observed in Opossum that at the moment of sperm pairing a gap junction arises between the two sperm membranes. B. SPERMATOZOA POSSESSING TWO AXONEMES

This is a characteristic feature of the whole group of orders known as Rhynchotoids (Fig. 15A, B); namely, the Psocoptera, Mallophaga, Anoplura, Thysanoptera and Rhynchota. In all instances, the doubling of this structure concerns only the axonemes and the centriole adjuncts, with the exception of. the mitochondria1 derivatives which are two in number (almost in .all cases) or even a single one (in Thysanoptera). Psocoptera generally have a single axoneme (Phillips, 1969); however, in one species, Atropos puZsatorium (Baccetti et al., 1969d), a high percentage of biflagellate sperms are found among the normal ones. It should be kept in mind, with regard to this, that among insects the accidental appearance of sperms with two axonemes is relatively frequent. This may be experimentally enhanced by mercaptoethanol treatment (Friedlander and Wahrman, 1965). It was reported by Baccetti et al. (1 969d) that, among Mallophaga, two equal axonemes consistently originate from two closely apposed centrioles lying beneath a rather enlarged nucleus. An analogous condition is found in Anoplura, among which Pediculus (Ito, 1966) and Haemutopinus (Baccetti et al., 1970a) have been studied. Among Thysanoptera (Fig. 15B) two axonemes are still encountered (Baccetti et al., 1969d); here, however, the accessory tubules, radial links and central sheath are wholly absent and the four central tubules, together with the 18 doublets, haphazardly form into a single bundle. It is curious that in such an arrangement Fig. 25. The origin of accessory tubules and accessory bodies in the spermatozoon of Bacillus rossius. (From Baccetti ef nl., 1972b.) (A) The tubules B of the doublets form a long outer arm, (a), and are surrounded by a system of Golgi (g) cisternae symmetrically disposed. ~ 6 0 , 0 0 0 .(B) The outer arms of the B tubules produce the accessory tubules (arrows). A Colgi (g) membrane is folded forming the precursor of an accessory body (b). ~180,000. (C) A later spermatid, with complete accessory tubules (at) and the rudiment of an accessory body (b) arising from a Colgi vesicle (g). ~90,000.(D)Migration of Cold vesicles (g) towards the axoneme. ~ 6 0 , 0 0 0 .

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the axoneme still retains its motility (unpublished observations on Thrips). Among Rhynchota the condition with two axonemes has the tendency to disappear. The heteropteran, Pyrrhocoris (Furieri, 1963b) has two axonemes which lie side by side, one longer than the other-a situation not found in other species. Other Heteroptera, e.g. Gelastocoris (Payne, 1966) and Nepa (Werner, 1966) have a single axoneme. The same is true of the most evolved Rhynchotoids, i.e. the Hornoptera, whose highest forms, the coccids, have non-flagellate spermatozoa. The presence of two axonemes, therefore, does not seem to be a haphazard event; it is a condition which distinguishes the primitive Rhynchotoid orders but disappears in the vast majority of the species in the most evolved orders. The type of motility acquired is rather peculiar and disorderly (see below). It is difficult to foresee what its selective advantage can be. C. NON-FIaAGELLATE SPERMATOZOA

This seemingly paradoxical condition appears, as far as I know, only in four insect groups (in one of which in two different forms). In three cases it appears to be associated with non-motility (the sperm being passively displaced), in the other two cases there are alternative organelles concerned with movement. Among the latter, the most widely known example is that of coccids mentioned above. In these Rhynchotes the syncytial condition which is shared by all insects during a given period of spermiogenesis (64 spermatids in Drosophila; 256 in Dacus-according to Baccetti and Bairati (1964); 128 in Atropos-according to Baccetti (1967)-and in Trychopteraaccording to Phillips (1970b); always increasing by the 2nd power) is retained by the adult sperms. In the light microscope Nur (1962) observed sperm bundles consisting of 16 elements in Pseudococcus obscurus, 32 in Eriococcus and Parlatoria, 64 in Puto. Moses and Coleman (1964) found in Steatococcus tuberculatus 32 spermatozoa in every bundle, each of them exhibiting a set of parallel microtubules surrounding the nucleus two and a half times. Robison (1966, 1970) and Ross and Robison (1969) have studied Parlatoria oleae, Pseudococcus obscurus, Matsucoccus bisetosus, Stomacoccus plantani, Kermes sp. and Put0 albicans. In the last species, spermatozoa generally exhibit a “cork-screw” shape. A motile apparatus, consisting of an alignment of 20-250 microtubules, spirals around a central chromatin axis (Fig. 16). Nuclear membrane,

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acrosome, mitochondria, centrioles and axoneme are all absent. The microtubules contain ATPase (Moses, 1966) and their arrangement always follows asymmetrical patterns. According to Robison (1 970) these features suggest that the microtubule bundle is associated with sperm motility. Ross (1 97 1) points out that the cork-screw shape is attained by virtue of a second microtubule system helically coiled around the spermatid, thereby moulding its characteristic outline. On the contrary, in one case in the Termites, replacement devices do not exist, hence the absence of a flagellum is accompanied by non-motility. In another instance replacement structures are available. The loss of flagella is progressively acquired in the course of evolution of the different families, the most primitive of which seem to possess normal flagellate spermatozoa. Some examples were succinctly described at the light microscopical level by Springhetti (1963). In the more primitive Mastotermitidae and Termopsidae, the spermatozoon is elongated and bears a flagellum; it is likely to be of the orthopteroid type encountered also in Embioptera (Baccetti et ul., 1972e). But in Kalotermitidae and Rhinotermitidae, spermatozoa appear to lack the flagellum. Baccetti et al. (1972e) have described this spermatozoon in a specimen from the first family, Kalotermes flavicollis and in one belonging to the second family, Reticulitermes Zucifugus. In the latter (Fig. 27) the spermatozoon is a severely involuted cell. It has a compact almost perfectly spherical shape. The acrosome is absent. At the posterior pole, a small cytoplasmic rim accommodates a few mitochondria of a conventional pattern, namely spheroidal in shape and small in size. Also present are two extremely short parallel cylinders looking like centrioles, but which are actually two truncated axoneme apices lacking the two central tubules and only provided with nine doublets at their periphery. These two flagellar anlagen do not protrude from the roundish sperm body which is made up solely of a head and unable to move. In Kulotermes (Fig. 28) a much more bizzarre shape is encountered. During spermiogenesis, the nucleus of each spermatid is condensed and flattened into an ovoid shape by a set of microtubules which surround it. The plasma membrane is separated from the nuclear membrane by a most tenuous interspace in which the microtubules are seen to run in a single array even in the mature sperm. The plasma membrane is displaced at four angles into four long arms which are also completely occupied by microtubules. Thus, apart from the acrosome, this sperm is devoid of flagellar anlagen and mitochondria. This situation closely resembles that

Fig. 26. Cross sections of mature sperms of Thelmatoscopus albipunctatus: a, acrosome; m, mitochondria1derivative; n, nucleus. The axoneme is lacking. ~120,000.(From Baccetti e t al., 1972c.)

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Fig. 21. The spermatozoon ofReticulitermes lucifugus.~ 6 0 , 0 0 0 .

Fig. 28. The spermatozoon of Kalotermes jlavicollis. ~ 3 9 , 5 0 0 .

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found in coccids. How this spermatozoon moves, if it does, is unknown. However, the long flattened arms crammed with microtubules suggest a pattern of undulating membranes. In the Diptera Psychodidae, Baccetti et al. ( 1 9 7 2 ~have ) examined several species, in which a common behaviour was pointed out. Their spermatozoon (Fig. 26) looks like a rather stiff long needle, with a multilayered cortical zone. This is actually the result of an association of the plasma membrane with the secretory products occurring in the testis lumen, which surround each germ cell. Inside this theca there is a nucleus which is highly elongated, compact and wholly or partly enveloped by a profuse material which is derived from the Golgi and may be viewed as an acrosome. Embedded in the acrosome is a small mitochondria1 derivative which has few cristae. These spermatozoa, which distinguish the whole Psychodidae family, are also completely immotile. In the genus of Protura, Ementornon, the spermatozoon has the form of a disk when contained in the testes. The circular nucleus is perispherical and flattened mitochondria lie in the central area. After ejaculation, the cell assumes the form of a cup. X. MOTILITY A. MOTILE MECHANISMS

The problem of insect sperm motility is centred around an understanding of ciliary motion of which it is a variation. Indeed, there are a series of variations of increasing complexity which parallel an increase in structural complexity of both supporting and contractile tail structures. Most attempts to explain ciliary or flagellar motion have so far focused on the flagellum of flagellate Protozoa or on the sperm tail of the sea urchin, both of which have many similarities. They both have a plain axoneme of the 9 + 2 pattern. Some experiments have been performed on the bull spermatozoon, whose axoneme is more complex ( 9 + 9 + 2) but whose general pattern agrees with the preceding model, being a sperm with a large head and a short tail. From the overall information thus far collected (Holwill, 1966; Brokaw et al., 1970; Brokaw, 1971) two conclusions are evident: (1) In Protozoa and in sea urchin sperm, motion consists of consecutive waves propagated along the tail thus resulting in a planar beat. Bull spermatozoa, however, show a helicoidal motion.

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(2) In spermatozoa these waves originate from a point situated in the centriole zone (Bradfield, 1955). This was demonstrated by amputations performed by Bishop and Hoffmann-Berling (1959), Goldstein and Brokaw (1968) and Goldstein (1 969) with a laser microbeam, and by Van Herpen and Rikmenspoel's (1969) indirect estimates concerning the size of this control centre on the basis of its susceptibility to X-rays and protons. In Flagellata, on the contrary, the amputated portion of the flagellum can beat and a different origin of waves must be supposed (Goldstein et al., 1970). At present, any explanation as to the role played by the single fibrillar components in motility and the mechanisms involved in wave triggering and propagation, are a matter of speculation. The presence of directly contractile structures has been debated. Gray (1 955) and Machin (1 958) considered that they were not associated with any precise flagellar component, whereas others have identified them with the two central tubules (InouC, 1959) or with the nine doublets which are the only universally occurring structures also having ATPase in their arms (Bradfield, 1955; Silvester and Holwill, 1965). Satir (1961) advanced a hypothesis based on the twocomponent mechanism of muscle contraction. He suggested that a myosin-like component might exist in the tubules and an actin-like one in the matrix, which for the first time is thus believed to be involved in flagellar motility. Along this line, again under the influence of muscular contraction, Satir (1968) and Brokaw (1968, 1971) put forward a new view on a sliding model, excluding the matrix and proposing (Satir) a sliding of each B subfibre with respect to the others. This fact cannot be demonstrated in sperm tails, where the orientation of the bent tip is difficult to determine. A separation between A and B subfibres has occasionally been described at the tip of the tail in Insect spermatozoa, but this does not demonstrate any form of sliding. Another hypothesis has been put forward by Schreiner ( 197 1) on purely mathematical grounds. He excludes contraction along any type of tubule and postulates the existence of welding points between the matrix and motile tubules along the flagellum, which would lead to the displacement of the straight segments during wave propagation. The active part of the movement would therefore be the recovery stroke. Nothing is known about this in the insect sperm flagellum where the recent demonstration of actin filaments, apparently independent of the microtubules (Behnke et al., 1971), suggests the presence of a muscle-like

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Fig. 29. Sequences of frames from high-speed cinefilms showing flagellar beat of different insect sperms. (A) Ctenocephulus mnis: 9 + 2 model. Total time: 0.21 s. x380. (B) Gryllotulpu gryllotulpu: 9 + 9 + 2 model with a rigid caudal tip (t). Total time: 0.43 s. x560. (C) Huematopinus suis: 9 + 9 + 2, the axoneme twinned in each sperm. Total time: 0.21 s. x790.

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mechanism. In connection with the helical shape of the waves, the structures which exhibit a helical arrangement around the sperm main axis may be thought to play an important role in the progression of the contractile events. These structures are the central sheath and the radial links. It is interesting to recall that the doublets run parallel to the sperm main axis. Motion pictures of sperms from different species show at least three forms of movement which seem to correspond to a structural hierarchy. The spermatozoa (9 + 2 axonemal pattern) of some species show a simple, almost planar wave of low frequency with a wave length as long as, or longer than, that of the flagellum. Tails are generally inextensible and a single wave type arising in the centriolar zone is propagated along them. The example examined by us is the spermatozoon of Aphaniptera (Fig. 29A). This has the simplest type of motility which closely resembles that of flagellates, marine sperms and cilia in general. At a second level of complexity there is a helical wave (owing t o two harmonic movements of equal amplitude). This model (Fig. 29B) is typical of all the species possessing the 9 + 9 + 2 formula, with accessory tubules and ATPase-rich accessory fibres. This type of movement and structure occurs in many insect and mammalian sperms. At the third level of complexity there is a combination of the two types of undulation. This is the form of movement which occurs in spermatozoa having a 9 + 9 + 2 tail plus accessory bodies endowed with ATPase activity. In some cases these bodies enable the sperm to course a doubly helical trajectory which remains straight (Tenebrio: Fig. 30A), in other cases their shape compels them to follow always doubly helicated trajectories in a helical course (Bacillus: Fig. 30B). A detailed study on sperm motility (Baccetti et al., 1972b) deals with this last form of movement which incorporates the preceding ones. The relevant deductions are as follows. Initiation of motility takes place in the centriolar region and has been identified in the axoneme apex which is supplied with laminae showing ATPase activity. This motor unit is called initiation motor (it must be remembered that the mature insect sperms lack a centriole). In this region it is postulated that consecutive small range revolving movements occur around the central axis, leading to related displacements between the bundle of tubule apices and the centriole adjunct. This torsional impulse propagates passively along the tail, through the assembly of tubules by virtue of their inextensibility. The internal viscosity of this system involves some energy expendi-

Fig. 30

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twe during the transmission of the movement. For this reason the first deformation stops almost at once and is of limited mechanical extension. This deformation, however, is enough to trigger a contraction in all the axonemal structures possessing ATPase activity (central sheath, arms, coarse fibres), possibly reacting with the actin units and thus propagating the contractile wave along the flagellum; this wave propagation is an active one. Tubules function only to propagate the mechanical deformation to consecutive levels. This wave is not reflected, but it dies away near the end of the flagellum. Upon it, at a following moment, a second wave of greater amplitude is superimposed. It arises when the oscillation frequency of the initiation motor nears the natural oscillation frequency of the structures which surround the axoneme. The resulting resonance enhances the oscillation amplitude of the outer structures until a mechanical deformation is also produced outside the axoneme, such as to evoke the active reactions of the extraaxanemal bodies which also are endowed with ATPase activity. The greater structural consistency of these organelles results in wave propagation towards the caudal end, as if it was an elastic wave which is reflected by the tail tip. This reflection occurs in a reverse direction, since it may be tentatively postulated that the surrounding medium acts as a boundary for the tail. The helicoidal trajectory is determined by the non-rectilinear shape and non-linear reactivity of the structure of the sperm tail. When the above observations are extended to more simpler cases, it may be deduced that where extraaxonemal bodies are not available a single cylindrical-helicoidal wave type arises (simple 9 + 9 + 2 insects). When the ATPase-containing coarse fibres lying between the accessory tubules are also absent, the quasi-planar wave type is evoked. This wave form has a much lower frequency, but a greater length being as long as the sperm tail and it is characteristic of the 9 + 2 model (first type of movement, as for instance among Aphaniptera). Other motility patterns, related to a particular flagellar architecture have emerged. As already mentioned, Gryllotalpa (Baccetti et al., 1971b) has a tail divided into two segments. The first has a Fig. 30. Sequences of frames from high-speed cinefilms of flagellar beat of different insect sperms. (A) Tenebrio molitor: 9 + 9 + 2 model with little accessory bodies: the trajectory is double helicated, but linear. h, Head. Total time: 0.17 s. x460. (B) BuciZlus rossius: 9 + 9 + 2 model with enormousaccessory bodies. The trajectory is double helicated and circular. h, Head. Total time: 0.09 s. x430.

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conventional 9 + 9 + 2 pattern, without accessory tubules and moves according to the general insect model. The second, with sclerified tubules, is completely rigid (Fig. 29B). The biflagellate Rhynchotoid sperm also moves in an irregular fashion. For instance, in Anoplura a rather ill-coordinated movement is found (Fig. 29C), although it belongs to the category of a single cylindrical helicoidal wave, short and frequent (general type), as suggested by the absence of accessory bodies. Movement of the complex syncytium of sperms in Coccids is most peculiar (Robison, 1970) and can possibly be regarded as helicoidal, like the motion of flagella (on which, however, the various wave types have not been studied). Since the accessory bodies are lacking, it should be ascribed to the two simpler categories, possibly to the second one. B. METABOLIC ASPECTS OF MOTION

In the preceding sections it was seen that in insect spermatozoa an important energy source is due to intense anaerobic glycolysis occurring in the axonemal complex. Normal respiration effected in the mitochondria1 compartment, when present, may be added to it. In all cases the stored polysaccharide is glycogen, which usually accumulates in the accessory tubules at the spermatid stage. The mature spermatozoon can thus call upon an endogenous store of carbohydrate. However, since a high carbohydrate level exists in both the seminal fluids and the spermatheca secretions, these may be thought of as being somehow a part of the material metabolized by the spermatozoon, despite the fact that the uptake of carbohydrate, or even indications of any type of pinocytosis, have never been demonstrated. Indeed, in many cases, sperm locomotion was observed even without carbohydrate administration (Hughes and Davey, 1969). In normal respiring sperms, oxygen is an element which regulates their motion. It may be experimentally demonstrated not only that these sperms are immotile under anaerobic conditions, but also that they migrate towards oxygen gradients. It was suggested that the tracheae-rich female conceptacula can thereby direct the movement of spermatozoa (Rao and Davis, 1960). Spermatozoa devoid of mitochondria are insensitive to oxygen (Baccetti e t al., 1972b). Hydrogen concentration also exerts an important effect on sperm motility: an optimum between pH 7.0 and 7.4 was demonstrated in Cimex by Rao and Davis (1 969). Richards (1963a) was the first to show that in Periplaneta sperm

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motility is a function of temperature and attains a maximum at 39". Davey (1958) has made similar observations. In Bacillus sperm (Baccetti et al., 1972b) motility was enhanced up to 50"C, but then it decreased though still remaining high up to 60°C. At this elevated temperature, however, few sperms were seen to survive. Temperature does not seem to appreciably influence wave length, but it does enhance both wave frequency and velocity. Motility is obviously affected by the osmotic pressure of the medium. In Periplaneta, Davey (1 958) reported a maximum activity with a tonicity exceeding 400 mosmoles. Later, lower tonicities were tested (Hughes and Davey, 1969). Generally, in flagella, altered conditions of viscosity affect the frequency, but not the form of beat (Brokaw, 1963, 1966; Holwill, 1965). Nevertheless, the high viscosity seems to reduce to two dimensions the normal threedimensional motion of bull spermatozoa (Rothschild, 1961). Studies on this problem in insect sperms are in progress. . C. THE PROBLEM OF SPERM CAPACITATION

This subject has been little investigated in insects. As observed by Hughes and Davey (1969) the general concept we can reach at present is that in some species fully mature sperms are transmitted to the female, while in other species their maturation is achieved in the female organism, thereby leading to a form of capacitation. In Periplanetu, for example, the extraacrosomal space is reduced in the spermatheca (Hughes and Davey, 1969). Likewise, in Sciuru a portion of the Nebenkern is eliminated (Makielski, 1966). In coccid insects the spermatozoa originally assembled under syncytial conditions become free individuals (Robbins, 1965). As stated earlier (Section 11), the most outstanding transformations occurring in the spermatheca concern the digestion and remoulding of the glycoprotein materials overlying the plasma membrane (Payne, 1934, in a hemipteran; Riemann, 1970; Riemann and Thorson, 1971; Phillips, 1971, in many Lepidoptera; Renieri and Vegni, 1972, in Iocusts). A form of capacitation may perhaps be recognized in these aspects, which in the moths Trichoplusiu and Anugustu culminates in the formation of an extracellular rod-like crystalline structure running parallel to the sperm, embedded in two thick crystallized sheaths. All this complex comes into being exclusively within the female genital tract and it originates from the material which formed the laminated appendages and the satellite body (Riemann, 1970; Riemann and

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Thorson, 197 1). The fine structure of the penetrating spermatozoon still requires more detailed study. XI. SPERMATOZOA POLYMORPHISM AND GENETICS

After the earliest work by von Siebold (1836), who discovered an atypical spermatogenesis associated with a typical one in a Mollusc, a number of investigations have been focused on the typical two-fold spermatogenesis found in many invertebrates; in particular in Lepidoptera where it was brought to light by Meves (1903). Fain-Maurel ( 1966) and Zylberberg ( 1969) have summarized its most relevant aspects. Among the various insect orders, when an atypical sperm line exists, it is generally a diploid or polyploid hyperpyrene line (Richards, 1963b; Bouix, 1963) because of variations in the ratio of DNA to nuclear proteins (Ansley, 1958). On the contrary, in Lepidoptera either an apyrene line is found when the nuclei are entirely eliminated during spermiogenesis, or an oligopyrene line when some nuclear residues persist (Zylberberg, 1969). Hyperpyrene spermatozoa have a normal length, but much thicker heads and tails as compared to normal sperms. They have been little studied as yet; the apyrene ones in Lepidoptera have been the object of recent research on the parts of Zylberberg and of Phillips (197 1). In these latter cells the nucleus does not undergo its typical condensation during the spermatid stage, as is true of eupyrene sperms; it remains spherical, glides caudal along the flagellum and then is eliminated. The acrosome does not form and the centriole comes to lie in the anterior cell portion. The corpuscle, interpreted by Zylberberg (1969) as a centriole, is conversely interpreted as a cap of undefined material by Phillips (1971). Both workers have found defective mitochondria1 derivatives in the apyrene line. In addition, Phillips (1 97 1) finds hollow instead of solid accessory tubules and, as already stressed, an absence of lacinate appendages around the plasma membrane. The presence or absence of the nucleus is not therefore related to the absence of any other organelle (excluding the acrosome, which forms on the nucleus itself), but rather is seen to affect normal organization in the spermatocyte stage. In fact, the cytoplasm has already received all its necessary information from the nucleus, and the spermatid nucleus is dormant (Olivieri and Olivieri, 1965). Many important papers on this aspect have appeared describing the long series of investigations on Drosophila mutants lacking the Y chromosome (Meyer, 1964, 1968; Kiefer, 1966;

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Anderson, 1967). Drosophila heydei does not progress beyond the spermatocyte I stage (Hess and Meyer, 1963); in Drosophila melanogaster, on the contrary, spermiogenesis begins without going to completion. All the spermatozoa of these X-0 males possess: ( 1) ill-organized mitochondria1 derivatives, sometimes supernumerary ones, with an anomalous crystalline core; (2) distorted or broken axonemes, which may lack some of its units; (3) bundles each having an overall smaller number of sperms which have shorter tails because they are immature. On the other hand, in the XYY males, spermatozoa are twice as long as normal ones (Meyer, 1968). Hence, in Drosophila. the Y chromosome seems to be endowed with a given number of fertility factors essential for the development of normal functioning spermatozoa. These factors must come into play during spermatocyte I stage since in normal X males spermatozoa are also normal. In this phase, similar loops to those of the lampbrush chromosomes are seen to form. In special mutants in which the Y chromosome is shorter (Bairati and Baccetti, 1966) and lacks one or more loops, sperms do not undergo maturation, thus resembling those from X-0 males. These individuals are sterile (Hess and Meyer, 1968). A particular mutant with a short Y (KL- 1-), studied by Kiefer (1969), has appgrently normal and motile spermatozoa and is capable of fertilizing the female. However, these spermatozoa degenerate within the female. They probably bear biochemical deficiencies which cannot be detected at the morphological level. In summary, the hypothesis is forwarded that the Y chromosome does not code for the proteins required in the formation of the sperm organelles, but governs the coordination of the various synthetic and morphogenetic steps (Meyer, 1970). Similar malformations have been induced by heat (Anderson, 1967) and X-rays (Hess, 1965). Altering the haemolymph composition by adding K + and Mg2+ (Meyer, 1969) also induces similar effects. All these altered features may in fact be viewed as phenocopies of Y deficiencies. What happens in the normally X-0 species (Orthoptera, Protenor) under the same experimental conditions has not been studied. Neither is it known whether a dimorphism between X-sperms and 0-sperms is detectable in those species in which a large X occurs, while their autosomes are minute. Almost all the genetics of spermatozoa still need studying.

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ACKNOWLEDGEMENTS

The original research reported in this article was supported by C.N.R. I am indebted to Drs. A. Bairati, jr., R. Dallai, R. Folliot, P. L. Maillet, M. Mazzini, E. Perotti, D. M. Phillips, S . Pratt and W. G. Robison for permission to use figures. REFERENCES Afzelius, B. (1959). Electron microscopy of the sperm tail. Results obtained with a new fixative. J. biophys. biochem. Cytol. 5,269-278. Allen, R. D. (1 968). A reinvestigation of cross-sections of cilia. J. Cell Biol. 37, 825-83 1. Anderson, W. A. (1 967). Cytodifferentiation of spermatozoa in Drosophila melanogaster. The effect of elevated temperature on spermiogenesis. Molec gen. Genet. 99, 257-273. Anderson, W. A. and AndrB, J . (1968). The extraction of some cell components with pronase and pepsin from thin sections of tissue embedded in an epon-araldite mixture. J. Microsc. 7,343-354. Anderson, W. A. and Ellis, R. A. (1967). Cytodifferentiation of the crayfish spermatozoon: acrosome formation, transformation of mitochondria and development of rnicrotubules. 2.Zellforsch. mikrosk. Anat. 77,80-94. Anderson, W. A. and Personne, E. (1 970). The localization of glycogen in the spermatozoa of various invertebrate and vertebrate species. J. Cell Biol. 44, 29-5 1. AndrB, J. (1 959). Etude au microscope electronique de l’evolution du chondriome pendant le spermatogknkse du Papillon du Chou Pieris brassicae. Annls Sci. nat. Zool. 12 Ser., 283-308. AndrC. J. (1961). Sur quelques ddtails nouvellement connus de l’ultrastructure des organites vibratiles. J . Ultrastruct. Res. 5 , 86-1 08. AndrB, J. (1962). Contribution 2 la connaissance du chondriome: Btude de ses modifications ultrastructurales pendant la spermatogBnkse. J. Ultrastruct. Res. Suppl. 3, 185 pp. AndrC, J. (1963). Some aspects of specialization in sperm. In “General Physiology of Cell Specialization” (G. Mazia and A. Tyler, eds), pp. 9 1-1 15. McGraw-Hill, New York. Andrt, J. (1965). A propos d’une leCon sur la Limule. Annls Sci. Clermon t-Ferrand 26, 2 7-3 8. Andrb, J. and Bernhard, W. (1 964). The centriole and the centriolar region. In “XIth International Congress Cell Biology”. Providence, R.I.9. AndrB, J. and ThiBry, J.-P. (1963). Mise en Bvidence d’une sous-structure fibrillaire dans les filaments axonkmatiques des flagelles. J. Microsc. 2, 7 1-80. Ansley, H. R. (1958). Histones of mitosis and meiosis in Loxa flavicollis (Hemiptera). J. biophys. biochem. Cytol., 4, 59-62. Austin, C. R. (1948). Function of hyaluronidase in fertilization. Nature, Lond. 162. 63-64.

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Rothschild, Lord (1961). Sperm energetics: an account of the work in progress. In “The Cell and the Organism” (J. A. Ramsay and V. B. Wigglesworth, eds), pp. 9-21. Cambridge University Press. Sakai, A. and Shigenaga, M. (1 967). Behavior of cytoplasmic membraneous structures in spermatogenesis of the grasshopper A tractomorpha bedeli Bolivar. Cytologia 32, 72-86. Sandoz, D. (1970). Evolution des structures au cours de la formation de l’acrosome du spermatozo’ide chez la souris. J. Microsc. 9, 535-558. Satir, P. (1961). “The mechanisms of ciliary motion: a correlation of morphe logical and physiological aspects of ciliary beat of the gill epithelium of the freshwater mussel, Eliptio complanatus, Solander” (Ph.D. Thesis). The Rockefeller Institute, New York. Satir, P. (1968). Studies on cilia. 111. Further studies on the cilium tip and a “sliding filament” model of ciliary motility. J. Cell Biol. 39, 77-94. Schreiner, K. (1971). Mechanics of ciliar and flagellar motion. Expl. Cell Res. 67,257. Shay, J. W. and Biesele, J. J. (1 968). Ultrastructural observations on spermiogenesis in the cave cricket Ceuthophilus secretus (Scudder). La Cellule 67, 269-282. Shay, J. W., Dobson, W. J., Simmons, E. E., Biesele, J. J. and Breland, 0. P. (1969). Subunits of flagellar accessory tubules. Tissue Cell 1, 593-596. Shelanski, M. L. and Taylor, E. W. (1968). Properties of the protein subunit of central-pair and outer-doublet microtubules of sea urchin flagella. J. Cell Biol. 38,304-3 15. Shoger, R. L. and Brown, G. G. (1970). Ultrastructural study of sperm-egg interactions of the horse crab Limulus polyphemus, L. (Merostomata: Xiphosura). J. submicrosc. Cytol. 2, 167-179. Shoup, J. R. (1967). Spermiogenesis in wild type and in a male sterility mutant of Drosophila melanogaster. J. Cell Biol. 32,663-675. Siebold, C. T. von (1836). Arch. Anat. Physiol. Wiss. Med. 232-255. Silvester, N. R. and Holwill, M. E. J. (1965). Molecular hypothesis of flagellar activity. Nature, Lond. 205,665-668. Springhetti, A. (1 963). Sulla struttura delle vescicole seminali delle termiti. A tti Accad. Naz. Ital. Ent. R c . 11,212-220. Stephens, R. E. (1970). Thermal fractionation of outer fibre doublet microtubules into A- and B- subfibre components: A- and B- tubulin. J. Molec. Biol. 47, 353-363. Stevens, R. E., Renaud, F. L. and Gibbons, I . R. (1967). Guanine nucleotide associated with the protein of the outer fibres of flagella and cilia. Science 156, 1606-1607. Sud, B. N. (1961a). The “chromatid body” in spermatogenesis. Q. J. Microsc. Sci. 102, 273-292. Sud, B. N. ( 196 1b). Morphological and histochemical studies of the chromatoid body and related elements in the spermatogenesis of the rat. Q. J. Microsc. Sci. 102,495-505. Tandler, B. and Moriber, L. G. (1965). Fine structure of the chromatoid body during spermatogenesis in the water-strider, Gerris remigis (Say.). 2. Zellforsch. mikrosk. Anat. 6 8 , 301-307. Tandler, B. and Moriber, L. G. ( 1966). Microtubular structures associated with the acrosome during spermiogenesis in the water-strider Gerris remigis (Say.). J. Ultrastruct. Res. 14, 391-404.

396

B. BACCETTI

Tates, A. D. ( 197 1). “Cytodifferentiation During Spermatogenesis in Drosophila melanogaster”. Drukkerij J . H. Pasmans, ~’Gravenhage. Taylor, J. H. (1 964). The arrangement of chromosomes in the mature sperm of the grasshopper. J. Cell Biol. 21, 286-289. Thomas, M. B. (1 970). Transitions between helical and protofibrillar configurations in doublet and singlet microtubules in spermatozoa of Stylochus zebra (Turbellaria, Polycladida). Biol. Bull. 138, 21 9-234. Van Herpen, G. and Rikmenspoel, R. (1969). Radiation damage done to bull sperm motility. 1. X-ray effects and target theory. Biophys. J. 9 , 822-832. Wada, S. K., Collier, J. R. and Dan, J. C . (1956). Studies on the acrosome. V. An egg membrane lysin from the acrosomes of Mytilus edulis spermatozoa. Expl, CellRes. 10, 168-180. Warner, F. D. (1970). New observations on flagellar fine structure. The relationship between matrix structure and the microtubule component of the axoneme. J. Cell Biol. 47, 159-182. Warner, F. D. (1971). Spermatid differentiation in the Blowfly Sarcophaga bullata with particular reference to flagellar morphogenesis. J. Ultrastruct. Res. 35,210-232. Werner, G. ( 1964). Untersuchungen iiber spermiogenesi beim Silberfischen, Lepisma saccharina L. 2. Zellforsch. mikrosk. Anat. 63, 880-912. Werner, G. (1 965). Untersuchungen uber die Spermiogenesi bien Sandlaufer, Cicindela campestris, L. Z. Zellforsch. mikrosk. Anat. 66, 255-275. Werner, G. (1966). Untersuchungen uber die Spermiogenese bei einem Lankiifer, Carabus catenulatus, Scop., und der Skorpion-Wasserwanze, Nepa rubra, L. 2. Zellforsch. mikrosk. Anat. 73, 576-599. Wilkes, A. and Lee, P. E. (1965). The ultrastructure of dimorphic spermatozoa in the hymenopteran Dahlbominus fuscipennis (Zett.) (Eulophidae). Can J. Genet. Cytol. 7, 609-6 19. Yasuzumi, G. (1956). Spermiogenesis in animals as revealed by electron microscopy. 111. Formation and submicroscopic structure of the tail flagellum of fish, insect, bird and Mammalia. Proc. 1st. Reg. Conf. Electron Microscopy in Asia and Oceania, Tokyo, pp. 25 1-256. Yasuzumi, G. and Ishida, H. (1957). Spermatogenesis in animals as revealed by electron microscopy. 11. Submicroscopic structure of developing spermatid nuclei of grasshopper. J. biophys. biochem. Cytol. 3, 663-668. Yasuzumi, G. and Lee, K. J. (1966). Spermatogenesis in animals as revealed by electron microscopy. XVI. The microtubular structure and sites of thiamine pyrophosphatase activity in premature sperm of the Japanese crayfish. 2. Zellforsch. mikrosk. Anat. 73, 384-404. Yasuzumi, G. and Oura, C. (1964). Spermatogenesis in animals as revealed by electron microscopy. XIII. Formation of a tubular structure and two bands in the developing spermatid of the silkworm, Bombyx mori Linn6. 2 . Zellforsch. mikrosk. Anat. 64, 210-226. Yasuzumi, G., Fujimura, W. and Ishida, H. (1958). Spermatogenesis in animals as revealed by electron microscopy. V. Spermatid differentiation of Drosophila and grasshopper. Expl. Cell Res. 14, 268-285. Yasuzumi, G., Sugioka, T., Tsubo, I., Yasuzumi, F. and Matano, Y. (1970). Spermatogenesis in animals as revealed by electron microscopy. XX. Relationship between chromatoid bodies and centriole adjunct in spermatids of grasshopper, Acrida lata. 2. Zellforsch. mikrosk. Anat. 110, 23 1-242.

INSECT SPERM CELLS

397

Zirwer, D., Buder, E., Schiilike, W. and Wetzel, R. (1970). Lineardichroitische untersuchungen zur DNS-Organisation in Spermienkoffen von Locusta migratoria, L. J. Cell Biol. 4 1 , 4 3 1-434. Zylberberg, L. (1969). Contribution 21 l'btude de la double spermatogenbe chez un Lbpidoptkre (Pieris brassicae L. Pieridae). Annls Sci. nut. 2001.Ser. 12, 1 1 , 569-626.


Author Index Numbers with an asterisk refer to pages on which references are listed at the end of the paper

B

A Abbott, N. J., 258, 293, 300, 303, 305 * Adams, J. B., 189, 209, 212, 214, 215, 250* Adams, R. T., 147, 181* Afzelius, B., 343, 349, 384* Agarwal, H. C., 72,78,80, loo*, 108* Akov, S., 56,60, 100* Alberici, M., 35,43* Allais, J. P., 74, 101* Allen, R. D., 343,384* Allen, R. R., 75, 101* Alvarez, R., 153, 155, 181* Alving, B. O., 277,306* Ames, A., 293, 306* Anders, F., 188, 211, 216, 220, 221, 250* Anderson, E., 324,386* Anderson, W. A., 330, 332, 350, 383, 384* And& J., 316, 318, 322, 332, 338, 341,348,349,350,355,357,384* Ansell,G. B.,69, 95, 96, 101* Ansley, H. R., 382,384* Aranda, L. C., 126, 128, 132, 158, 160, 161, 178*, 180* Argarwal, H. C., 72, 78, 80, loo*, 108* Ariens, E. J., 11,46* Arnaiz, G. R. D., 35,43* Ashhurst, D. E., 264, 274, 293, 306* Asperen, K. Van., 276,306* Aurbach, G. D., 21,38,43* Ausit, E. G., 153, 155, 181* Austin, C. R., 326, 384*

Babad, H., 16, 37,42* Baccetti, B., 316, 317, 318, 319,320, 322, 324, 325, 326,327,328,329, 330, 331, 333, 334, 335,336,338, 339, 341, 242, 243,345,347,348, 349, 350, 351, 352,353,354,355, 357, 358, 360, 361, 362,363,364, 365, 366, 367, 369,370,371,372, 374, 377, 379, 380, 381, 383, 385*, 386*, 394* Bairati, A., Jr., 327, 338, 341, 345, 350, 351, 362, 370, 383, 385*, 386* Balasubramanian, A., 232, 233, 234, 246,250* Ballowitz, E., 367,386* Baptist, B. A., 231, 234, 235, 237, 238,246,250* Barker, K. R., 327,386* Barkley, D. S., 34,45* Barlow, J. S., 57, 105* Barnum, C. P.,100,107* Barron, E. S. G., 54, 108* Baumann,C. A.,57,61,89, 107* Bawa, S. R., 329, 349, 351,367,369, 386 Baxter, C., 151, 178* Bdolah, A., 15, 16,37,42* Beams, H. W., 324,386* Beaudoin, A. R., 72, 78, 83, 89, 90, 101 * Beck, S. D., 57, 101*, 188, 209, 210, 212,215,251* Behnke, O., 338, 341, 345, 347, 375, 386*

399

400

AUTHOR INDEX

Bentley, P. J., 39,40,42* Ben-Zvi, R., 16, 37,42* Berendes, H. D., 32,45* Bergerard, J., 74, 101* Berlin, L. C., 209, 210, 230, 231, 250* Bernard, R., 56,60, 106* Bernard, W., 332, 384* Bernheim, M. L. C., 54, 105* Bernini, F., 328, 329, 354, 371, 386* Berridge, M. J., 2, 3, 5, 7, 12, 13, 15, 16, 18, 19, 22, 23, 24, 25, 26, 27, 28, 29, 31, 42*, 46*, 47*, 267, 306* Bhamburkar, M. W.,75, 107* Bhat, J. V., 73,78, 109* Bieber, L. L., 55, 56, 60, 62, 65, 68, 70, 73, 78, 80, 81, 84, 85, 87, 88, 90,91, 101*, 107* Biesele, J. J., 324, 332, 334, 338, 348, 387*, 388*, 395* Biggers, J. D., 367, 386* Bigliardi, E., 318, 319, 320,333,334, 342, 347, 348, 351,352,357, 362, 365,369,385*, 386* Binstock, L., 286,306* Birge, W. J., 350, 386* Birt, L. M., 73, 76, 103* Bisho, D. W.,375, 386* Bjork, L. B., 329,389* Blakley, R. L.,54, 101* Blewett, M., 55, 56, 57, 101*, 104* Bloch, D. P., 33 1,386*, 388* Bloom, F. E., 35,47* Bloom, W.,37,42* Boistel, J., 63, 101*, 164, 178*, 259, 275, 277, 278, 281, 282,288, 289, 304,306*, 308*. 309* Bongers, J., i95, 2b2, 203, 204, 206, 250* Bonner, J. T., 34,45* Borkenhagen, L. F., 54, 101* Borle, A. B., 38,42* Bouix, G., 331,381,387* Boulton, P. S., 260,280, 306* Bourguet, J., 39,40,42*, 45* Bowen, R. H., 354,387* Boyme, T., 38,44* Brack, S. D., 331, 386*

Bracken, C. K., 77, 101* Bradfield, J. R. C., 329, 374, 375, 387*, 389* Brading, A. F., 35,42* Brady, J., 275,306* Brady, R. O., 54, 106* Branson, C. M., 37,45* Braun, G., 3,46* Breckenridge, B. McL., 34,42* Breland, 0. P., 324, 332, 334, 338, 348,387*, 388*, 395* Bremer, J., 54, 89, 102* Breucker, H., 317, 385*, 390* Bridges, P. M., 68, 109" Bridges, R. G., 57, 58, 61, 65, 66,61, 68, 69, 73, 74, 75, 76, 77, 78, 79, 80, 81, 83, 84, 85, 87, 88, 90, 91, 92, 93, 94, 97, 98, 102*, 103* Brighman, M. W.,302,303,306* Brodie, B. B., 38,45* Brokaw, C. J., 374, 375, 381, 387*, 390* Bronskill, J. F., 234, 238, 250* Brookes, V. J., 72, 73, 78, 81, 91, 101*, 106* Brown, A. W. A., 56, 58,72,78, 104*, 108*

Brown, B. M., 172, 173, 174, 175, 178* Brown, G. G., 316,387*, 395* B r u t , M., 57, 102* Bryan, J., 341, 387* Buck, J. B., 276,306* Buder, E., 332, 397* Bueding, E., 35,43* Biilbnng, E., 35,42*, 43* Burgos, R., 37,48* Bum, J. H., 34,42* Burrini, A. G., 316, 322, 324, 325, 326, 327, 329, 330,331, 333,334, 335, 336, 342, 343,345,341,348, 349, 350, 351, 352,353,355,357, 358, 360, 361, 362,363,364,365, 366, 361, 369, 372,374,317,379, 380,381,385*, 386* Biisgen, M., 185,250* Butcher, R. W.,12, 15,31, 35,36,37, 38,39,43*, 44*, 47* Byczkowska-Smyk, W.,338,391*

40 1

AUTHOR INDEX

Bygrave, F. L., 3 1,43*

C Cameron, I. R., 302, 306* Cameron, M. L., 348, 387* Cantacuzsne, A. M., 334, 335, 387* Cantoni, G. L., 54, 102*, 106* Caraboeuf, E., 277, 278, 306 Carasso, N., 40, 45* Carlson, A. D., 259,3 11 * Carnay, L., 176, 181* Carpenter, D. O., 277, 306* Carter, W., 217, 223, 250* Castillbn, M. P., 72, 102* Catalh, R. E., 72, 102* Chan, S. K., 76, 102* Chandler, W. K., 286, 306* Chapman, J. A., 264, 306* Chase, L. R., 21, 38,43* Chasey, D., 342, 387* Cheldelin, V. H., 57, 61, 62, 65, 73, 78, 80, 81, 84, 91, 101*, 105*, 107* Chevailler, P., 328, 329, 332, 387* chi, Y. Y., 34,43* Choi. J. K.., 39.40.43* , Chojnacki, T., 66; 73, 85, 96, 102*, 103* Clarke, R. G., 208, 241, 251* Clausen, T., 38,44, Claypool, C. J., 331, 388* Clayton, B. P., 324, 388* Clermont, Y., 334, 388* Cmelik, S. H. W., 72, 103* Coggeshall, R. E., 303, 306* Cohen,M. J., 119, 122, 128, 129, 130, 132, 133, 134, 135, 136, 138, 139, 142, 149, 150, 178*, 179*, 180* Coleman, J. R., 370, 392* Coles, M., 55, 56, 104* Colhoun, E. H., 63, 66, 103* Collier, J. R., 326, 396* Conrad, R. G., 37,45* Cook, E. F., 132, 181* Cooper, D. P., 317,394* Cooper, M. I., 56, 59, 103*

Cordle, M., 174, 179* Corning, W. C., 119, 126, 127, 128, 142,144,179* Costa, E., 34,48* Costin, N. M., 264,306* Cox, J. T., 57, 61, 65, 80, 91, 98, 102* Craig, A. B., 38,43* Creed, R. F. S., 367, 386* Crone, H. D., 73, 74, 75, 76, 78, 81, 84, 85, 86, 87, 103* Curan, M. M., 40,43*

D Dadd, R. H., 57, 103* Daems, W. Th.,351,388* Dallai, R., 316, 317, 322, 324, 325, 326, 327, 328, 329, 330, 331, 333, 334, 335, 336, 338,339,341, 342, 343, 345, 347, 348,349,350,351, 352, 353, 354, 355,357,358,360 361, 362, 363, 364, 365,366, 367, 369, 371, 372, 374, 377,379, 380, 381,385*, 386*, 388*, 394* Dan, J. C., 326,388*, 396* Danilova, L. V., 341, 388* Daoust, R., 334,388* Das, C. C., 331, 388* Dass, C. M., 329,388* Dauterman, W. C., 61, 65, 67, 68,69, 70, 80, 81, 84, 85, 86, 87, 89, 98, 103*, 105*, 107* Davey, K. G., 32, 43*, 380, 381, 388*, 390* Davies, R. G., 232, 233, 234, 246, 250* Davis, N. T., 215, 242, 251*, 380, 393* Davis, R. E., 220,251*, 255* Dawson, R. M. C., 54,84, 103* D’Costa, M. A., 73,76, 103* DeLong, D. M., 56, 106* Dempsey, P., 38,45* De Robertis, E., 35, 43* Deutsch, K., 324,388* Devine, R. L., 324,386*

402

AUTHOR INDEX

Diamond, J. M., 267,285,307* Dibella, F., 34, 46* Di Bona, D. R., 40,43* Dickerson, G., 193,252* Diecke, F. P. J., 259, 262, 278, 279, 281,282,293,312* Dingman, W.,168, 179* Disterhoft, J. F., 119, 126, 127, 128, 142, 144, 179* Djahanparwar, B., 164, 181 * Dobbs, J. W., 35,43* Dobson, W.J., 348,395* Dominas, H., 73,90, 110* Donald, R. A., 37,44* Doolin, P. F., 350,386* Douglas, W.W.,37,43* Drew, M. E., 209,250* Drummond, G. I., 17, 18,21,43* Duchlteau, G., 275,276,307* Dumm, M. E., 75, 107* Duncan, L., 2 1 , 4 3* Dupont, P., 59, 104* Duspiva, F., 21 6, 25 1*

Epstein, S . E., 16, 20, 36, 43*, 44*, 48* Ericson, L. E., 89, 104* Erk, F. C., 56, 104* Esch, I. Van., 276, 306* Etienne, J., 74, 101*

F

Fain-Maurel, M. A., 382,388* Farese, R. V.,49,49* Fast, P. G., 71, 72, 73, 74, 77, 78, 104* Favard, P.,40,45* Fawcett, D. W., 37, 42*, 303, 306*, 332,334,350,355,388*, 389* Fedak, S. A., 21,43* Feir, D., 188, 209, 210, 212, 215, 251* Ferraguti, M., 329,389* Fielding, L., 54, 101* Filshie, B. K., 194,255* Finder, A. G., 38,44* Fine, R. E., 341,389* E Fischer, R. W.,99, 108* Fisher, R. W.,259,308* Eddy, E. M., 334,388*, 389* Fitz, I. B., 59, 104* Eddleman, C. D., 324,388* Fleischer, N., 37,44* Edelman, I. S., 39,47* Florkin, M., 275,276, 307* Edwards, J. S., 188, 204, 205, 210, Folliot, R., 327, 345, 355, 364, 365, 238,25 1* 389* Forbes, A. R., 200, 220, 242, 243, Ehrhardt, P., 189, 191,252* Ehrlich, P., 257, 307* 244,25 1*, 254* Eisenstein, E. M., 115, 117, 119, 120, Forer, A., 338, 341, 345, 347, 375, 386* 121, 122, 123, 128, 129, 130, 132, 133, 134, 135, 136, 138, 139, 142, Fowler, K. S., 99, 106* 145, 149, 150, 155,165,168,169, Fraenkel, G., 55, 56, 57, 58, 59, 60, 170, 172, 174, 175, 178*, 179*, 61, 101*, 102*, 103*, 104*, 107*, 180*, 181* 108* Eldefrawi, M. E., 95, 103*, 266, 293, Francis, D., 34,43* 298,307* Fratello, B., 316, 324, 327, 338, 342, Eliakova, G. V., 330,388* 343,386* Elliot, A. B., 39,43* Frazer, A., 36,47* Ellis, J., 56, 105* Frazier, H. S., 40,44* Ellis, R. A., 330,384* French, E. W., 59, 104* Emmersen, J., 345, 375, 386* Friedlander, M., 332, 369, 389* Entman, M. L., 36,43* Friedman, N., 38,44*

403

AUTHOR INDEX

Friedman, S., 60,61, 104* Friend, W. G., 57, 104*, 188, 193, 196, 203, 206, 208,234,238,238, 250*, 251*, 255* Fuge, H., 341,389* Fujimura, W., 322, 396* Furieri, P., 328, 370, 389*

Greengard, P., 35,46*, 49,49* Grimstone, A. V., 336,343,349,389* Grzelak, K., 66, 105* Guillemin, R., 37, 48* Guinness, F. E., 40,46* Gupta, B. L., 267, 306* Guthrie, D. M., 132, 179* Gyrisco, G. G., 195,254*

G Gall, J. G., 329, 341, 389* Gardiner, B. 0. C., 11, 12, 33, 46* Carey, F. G., 67, 70, 85, 88, 104*, 110* Gassner 111, G., 332, 334, 338, 387*, 389* Gatenby, J. B., 324,325, 334, 389* Gay, H., 33 1,388* Geer, B. W.,56, 58,61, 62, 65,81,92, 99, 104* Gelber, B., 176, 179* Geschwind, I. I., 3 7 , 4 7 * Gibbons, I. R., 329, 336, 341, 343, 349,389*, 390*, 394*, 395* Gilbert, L. I., 75, 76, 109* Gingrich, R. E., 56,60, 104* Girardier, L., 38,44* Giusti, F., 324, 325, 327, 328, 329, 336, 338, 339, 343,348,349, 350, 351, 354, 357, 358,361,362,363, 364,365,371,385*, 386* Glassman, E., 174, 179* Goldberg, A. L., 34,44*, 48* Goldberg, M., 245, 25 1* Goldstein, S. F., 374, 375, 390* Goodchild, A. J. P., 191, 192, 207, 208,235,236,25 1 * Gordon, H. I., 89, 105* Gordon, R. M., 193,252* Gordon, S. A., 223, 25 1 * Goriachkina, V.,330, 388* Gosbee, J. L., 37,45* Gouranton, J., 33 1,392* Graves, J. B., 75, 106* Gray, J., 375, 390* Greenberg, D. M., 54,89, 102* Greenberg, M. J., 11,44*

H Habibulla, A. M., 86, 87, 105* Hackman, R. H., 245,25 1* Haeusler, G., 36,48* Haggerty, R., 119, 142, 179* Hagiwara, S . , 278, 307* Hagley, E. A. C., 216, 251* Hales, C. N.,'16,47* Halstead, W.C., 128, 179* Hamann, K. F., 3 , 4 6 * Handler, J. S., 39, 44* Handler, P., 54, 105* Hardman, J. G., 17,44* Harris, J. T., 124, 125, 127, 144, 180* Harris, P., 77, 101* Hashimoto, A., 73, 105* Hause, L. L., 304, 307* Hawkins, J., 35,43* Hawthorne, J. N., 69, 96, 101* Haylett, D. G., 38, 44* Hays, R. M., 40,44* Hazan, A., 58, 109* Hebb, C., 54, 105* Heim, F., 35,46* Hellyer, G. C., 68, 109* Henderson, A., 174, 179* Henion, W. F., 15, 16,44*, 47* Henley, C., 390* Hennig; E.., 189, 194, 195, 196, 207, 251* Herold, F., 327, 365, 390* Hess, A., 264,307* Hess, M. E., 36,47* Hess, O., 383,390* Hibbs, E. J., 209,210,230, 231, 250* Highnam, K. C., 3 2 , 3 3 , 4 4 * Hill, A. V.,294,307*

404

AUTHOR INDEX

Hill, L., 32, 33,44* Hinton,T., 56, 60, 61, 104*, 105* Hirumi, H., 244,25 1* Ho, R. J., 1 5 , 3 8 , 4 3 * Hoage, T. R., 324, 332, 390* Hodgkin, A. L., 94, 105*, 277, 291, 307* Hodgson, E., 57, 61, 65, 67, 68, 69, 70, 73, 74, 76, 78, 80, 81, 82, 83, 84, 85, 86, 87, 88, 89, 90, 91, 96, 98, 101*, 105*, 106*, 107*, 108*, 109* Hoffer, B. J., 35,47* Hoffman-Berling, H., 375,386* Hokin, L. E., 37,44* Holden, J. S., 87, 102* Holwill, M. E. J., 336, 374, 375, 381, 390*, 395* Honig, C. R., 38,43* Hopkins, J. M., 342, 347, 350, 390* Hori, K., 189, 195, 196, 207, 209, 210,215,238,241,251*, 252* Horie,Y.,57,58,61,78, 105* Horn, G., 156, 157, 180* Horowitz, B. A., 38,44* Horowitz, J. M., Jr., 38, 44* Horridge, G. A., 118, 124, 132, 134, 135, 142, 144, 145, 151, 152, 164, 180* Horstmann, E., 316, 317, 387*, 390* Hoshiko, T., 308* House, H. L., 57, 105* Houx, N. W. H., 80,100* Howson, D., 122, 127, 132, 142, 180* Hoyle, G., 122, 132, 144, 157, 158, 159, 180*, 250,275,281,307* Hubbard, J. I., 35,45* Hubbell, W.L., 176, 180* Hughes, G. M., 155, 180* Hughes, M., 380,381,390* Hughes-Schrader, S., 332,390* Hurrell, D. P., 72, 103* Hynie, S., 40,45*

I Idelman, S., 367,390* Imura, H., 15,45*

InouB, S., 332,375,391 * Ishida, H., 322, 329, 396* Ito, S., 322, 324, 369,391 * Ito, T., 57, 58, 61, 78, 105*

J Jacklet, J. W., 134, 135, 149, 150, 178*, 180* Jard, S., 40,45 * Jenkinson, D. H., 34, 38,44*, 45* Johnson, H., 149, 180* Jones, S. F., 35,45* Jordon-Luke, B. M., 324,388*

K Kafka, M. C., 40,45* Kakiuchi, S., 34,45* Kalra R. L., 7 2 , 7 8 , 8 2 , 105* Kamienski, F. X., 7 2 , 8 1, 106* Kamin, L. J., 127, 128, 180* Kandel, E. R., 151, 156, 157, 162, 163,164, 180* Kanfer, J. N., 54, 106* Kastings, R., 57, 106* Katz, B., 291,307* Kaufman, B. P., 33 1,388* Kaye, J. S., 324, 325, 326, 331, 334, 342,345,348,391* Kemp, P., 84, 103* Kennedy, E. P., 54, 101*, 106*, 109' Kerkut,G. A.,63, 106*, 176, 180* Kessel, R. G., 318, 324, 329, 332, 342,390*, 391* Keynes, R. D., 269,285,307* Khan, M. A. Q., 6 1 , 6 5 , 7 6 , 8 0 , 8 3 , 8 4 , 88,98, 105*, 106* Kiefer, B. I., 382, 383, 391* Kinsella, J. E., 73, 74, 106* Kinsey, M. G., 185, 187, 195, 196, 197,200,201,202,205,246,252* Klee, W.A., 54, 106* Kleeman, C. R., 302,306 Klein, J. R., 54, 105*

405

AUTHOR INDEX

Kloft, W., 189, 190, 191, 200, 201, 210,211,215,220,252* Koefoed-Johnsen, V., 286, 307* Konijn, T. M., 34,45* Kornberg, A., 54, 109* Korzybski, T., 66, 73, 85,96, 102* Kraicer, J., 37,45* Krasilovsky, G. H., 134, 179* Krieger, D. L., 57, 103* Krishna, G., 38,45 * Krnjevik, K., 34,45* Kropf, R. B., 67,70, 110* Kruitwagon, E. C., 204,208,255* Krzysztofowicz, A., 338, 391 * Kuffler, S. W., 258, 273, 303, 307*, 308 * Kulka, R. G., 18,37,45* Kumar, S. S., 65, 67, 68, 70, 85, 86, 87,88,90,101*, 106*, 107* Kunkel, H., 189,252* Kuo, J. F., 49,49*

Leenders, H. J., 32,45* Le Menn, R., 329,391 * Lemonde, A., 56, 60, 72, 78, 83, 89, 90, 101*, 106* Lenartowicz, E., 67,68, 106* Leste, R. L., 76, 102* Levey, G. S., 16, 20, 36, 43*, 44*, 48 * Lewis, S. E., 99, 106* Lilly, J. H., 57, 101* Lindley, B. D., 308* Lindwell, J. O., 94, 108* Lipsitz, E. Y., 74, 78, 106* Lipson, L.C., 40,45* Locke, M., 239,245,252* Luco, J. V., 126, 128, 132, 158, 160, 161,178*, 180* Lunat, M., 72, 103*

M L Lambremont, E. N., 74, 106* Landau, E. M., 35,45* Lane, N. J., 95, 109*, 258, 259, 260, 262, 264, 266, 267, 268, 270, 271, 272, 273, 274, 276, 277,278, 279, 280, 281, 282, 283,285,288,289, 290, 291, 296, 303, 307*, 308*, 309*, 311* Langslet, A., 36, 45* Lanzavecchia, C., 329,389* Lassota, Z., 66, 105* Laszlo, S., 60,61, 104* Laurema, S., 209, 210, 214, 215, 252*, 254* Lavanchy, P., 193,224,254* Lavoipierre, M. M. J., 252* Lea, A. O., 56, 106* Leaf, A., 40, 43* Lecar, L., 286, 306* Leclercq, J., 275, 276,307* Ledbetter, M. C., 341, 391 * Lee, K. J., 330, 396* Lee, P. E., 328, 396*

Machin, K. E., 375,391* MacInnes, J. W.,332, 391* Maddrell, S. H. P., 11, 12, 33, 46*, 259, 260, 261, 264,266,267,268, 271, 277, 278, 281,282,285,288, 303,304,308*, 31 1* Magis, N., 59, 107* Maillet, P. L., 327, 331, 364, 365, 389*, 394* Makielski, S. K., 338, 367, 381, 392* Malamed, S., 40,44* Maltbie, M., 2 1, 36,46* Mangos, J. A., 3,46* Mansingh, A., 63,66,84, 108* Maramorosch, K., 244,25 1 * Marchbanks, R. M., 94, 107* Martin, K., 94, 105* Mason, J. W.,34,46 Matano, Y., 334,396* Matschinsky, F. M., 34,42* Matsukura, S., 15,45* Matsuyama, H., 15,45* Matingley, R. F., 304, 307* Matty, A. S., 40,46* Mayer, S. E., 21,36,46*

406

AUTHOR INDEX

Mazzini, M., 324, 325, 327, 336, 343, 348, 350, 351, 357, 358,361,362, 364,365,385*, 392* McAfee, D. A., 35,46* McAllan, J. W., 189, 209, 212, 214, 215,250* McConnell, H. M., 176, 180* McFarlane, J. E., 57, 74, 78, 106*, 108* McGaugh, J. L., 169, 180* McGinnis, A. J., 57, 106*, 107* McIlwain, H., 34,45* McLean, D. L., 185, 186, 187, 195, 196, 197, 200, 201, 202, 205, 246, 252*, 253* McMaster-Kaye, R., 331, 334, 391 * Mehendale, H. M., 61, 65, 67, 68, 69, 70, 80, 81, 85, 86, 87, 89, 98, 105*, 107* Mehrotra, K. N., 63, 75,84, 98, 103*, 107* Meinertz, T., 36, 46* Mellon, D., 259, 31 1* Meng, H. C., 15,38,43* Menon, T., 34,48* Metz, C. B., 316,387* Metz, C. W., 367, 392* Meves, F., 33 1, 382, 392* Meves, K., 286, 306* Meyer, G. F., 357, 382, 383, 390*, 392* Meythaler, B., 35,46* Mezei, C., 85, 87, 103*, 107* Miledi, R., 34,45* Miles, P. W., 188, 189, 194, 195, 196, 197, 200, 202, 203,205,206,207, 208, 209, 210, 211,212, 215,216, 217, 220, 221, 223,224,232,234, 236, 237, 238, 239,240, 241,244, 245,247,249,253* Miller,P. L., 117, 181* Miller, R. L.,374, 387* Milligan, J. V., 37,45* Mitchell, H. K., 72, 110* Mittler, T. E., 57, 103*, 185, 197, 200,205,206,253* Mitznegg, P., 35,46* Miyake, T., 15,45 * Moericke, V., 197, 200, 205, 226, 227,228,239,244,253*

Mohri, H., 341,392" Monroe, R. E., 55, 56, 6 0 , 6 2 , 7 8 , 8 1 , 101* Moon, H. M., 174, 179* Moore, W.,56, 107* Morel, F., 39,42* Moreton, R. B., 95, 108*, 109*, 258, 259, 264, 266, 268,272,274,281, 282, 283, 284, 285,286,287, 289, 290, 291, 296, 300, 304, 308*, 309*,311* Morgan, M. E., 15, 36,47* Moriber, L. G., 327,334,395* Morris, D., 54, 105* Moses, M. J., 363, 370, 371, 392* Moskowit, J., 38,45* Moulton, B., 65, 68,87, 90, 107* Mukai, K., 73, 105* Municio, A. M., 72, 102* Munz, K., 327,365,390* Murphy, R. K., 145, 181* N Nagata, N., 14,21,38,39,46* Nakajima, S., 278, 307* Namm, D. H., 21,36,46* Narahashi, T., 258, 277, 278, 289, 308*, 312* Nault, L. R., 195,254* Nebel, B. R., 329,392* Nelson, L., 341,392*, 393* Nelson, W.L.,100, 107* Nesbett, F. B., 293, 306* Newburgh, R. W., 57, 61, 62, 65, 72, 73, 75, 78, 80, 81, 84, 85, 86, 87, 90, 91, 101*, 103*, 105*, 106*, 107* Nicholls, J. G., 258,303,308* Niemierko, S., 67, 68, 106* Noble,E.P., 172, 173, 174, 175, 178* Noland, J. L., 57, 60, 61, 89, 104*, 107* Noyes, D. T., 56, 105* Nuorteva, P., 209, 210,214,215,216, 224,232,233,252*, 254* Nur, U., 370,392* Nurnberger, J., 1 19, 126, 127, 128, 144, 179*

AUTHOR INDEX

0

407

Petri, L., 185, 254* Petrinovich, L. F., 169, 180*, 181* O'Brien, R. D., 95, 103*, 258, 266, Phillips, D. M., 318, 320, 322, 325, 293,298,307*, 308* 326, 327, 328, 332,334,338, 339, Offermeier, J., 11, 46* 340, 341, 342, 343,345,347,348, Olander, R. M., 58,92, 104* 355, 357, 358, 360,365,367,369, Oliver, A. P., 35,47* 370,381,382,389*, 383* Oliver, G., 176, 180* Piantelli, F., 322, 326, 327, 329,330, Olivieri, A., 382, 392* 331, 335, 336, 342, 345, 348, 349, Olivieri, G., 382, 392* 350, 351, 352, 353,360, 363, 365, Orloff, J., 39,44*, 46* 366, 367, 369, 377,379,380,381, Oschman, J. L., 3, 22,46* 385* Ossiannilsson, F., 244, 254* Pichon, Y., 95, 107*, 108*, 109*, Oura, C., 322, 348, 396* 259, 264, 267, 268,271,272,273, (Bye, I., 15, 36, 45*, 47* 274, 275, 276, 277,278,280,281, 282, 283, 284, 285,286,287,288, 290, 291, 293, 294,295,296,297, 299, 301, 304, 308*, 309*, 31 1* P Piechowska, M. J., 85, 103* Pilcher, D. E. M., 11, 12,33,46* Pak, C. Y.C., 40,45* Pilet, P. E., 224, 254* Paleg, L. G., 223,25 1* Pipa, R. L., 132, 181* Pallini, V., 322, 326, 327, 329, 330, Pitman, R. M., 63, 106* 331, 335, 336, 342, 345, 347, 348, Plowman, K. M., 341,393* 349, 350, 351, 352,353,357,360, Pochon-Masson, J., 330,331,393* 362, 363, 365, 366,367,369,377, Poisner, A. M., 37,43* 379,380,381,385* Polonovski, J., 74, 101* Pant, N. C., 55, 56, 107* Porter, K. R., 341, 391* Park, C. R., 38,44* Posternak, T., 15, 16,44*, 47" Pastan, I., 15,46* Potter, D. D., 273, 303, 307* Patel, N. G., 2,42* Powell, C. A., 17, 18,43* Patterson, E. K., 75, 107* Pratt, S. A., 355,357, 393* Pattillo, R. A., 304,307* Prendergast, J. G., 226, 231, 246, Patton, R. L., 57, 75, 81, 104*, 109* 254* Payne, F., 327,370,381,392* Price, G. M., 73, 75, 76, 79, 83, 94, Peachey, L. D., 40,46* 102* Peake, G. T., 37,48* Prince, W. T., 2, 3, 12, 13, 19,22,23, Pease, D. C., 341,392* 2 4 , 2 5 , 2 6 , 2 7 , 2 8 , 2 9 , 3 1 , 4 2 * ,47* Periti, P., 322, 326, 327, 329, 330, Pringle, J. W. S., 132,181* 331, 335, 336, 342,345,348,349, Pritchatt, D., 122, 132, 134, 142, 144, 350, 351, 352, 353,360,363,365, 145, 181* 366; 367; 369; 377; 379; 380,381, Pumphrey, R. J., 153, 154, 155, 157, 385* 164,181* Perotti, M. E., 327, 333, 335, 336, 341, 342, 343, 345,347,350,362, 386*, 392*, 393* R Perry, M. C., 16,47* Persijn, J. P., 351,388* Raine, J., 220,242,243,244,254* Personne, E., 350,384* Rall,T. W., 12, 31,34,45*, 48* Petersen, M. J., 39,47* Ramsay, J. A., 3,47*

408

AUTHOR INDEX

Ramwell, P. W., 40,47* Rad, H. V., 380,393* Rao, K. 0. P.,72,78, 108* Rao, R. H., 88, 108* Rasmussen, H., 2, 12, 13, 14, 19, 20, 21, 25, 27, 31, 34, 36, 38, 39, 40, 41,44*, 46*, 47* Rasso, S. C., 58, 108* Ratner, A., 37,47* Rawdon-Smith, A. F., 153, 154, 155, 157,164, 181* Redina, G., 54, 108* Reger, J. F., 317,393* 394* Reese, T. S., 302, 303,306* Reiser, R., 57, 109* Renaud, F. L., 341, 343, 394*, 395* Renieri, T., 324, 325, 336, 343, 348, 350, 351, 357, 358,361,362,364, 365,381,385*, 394* Retzius, M. G., 355, 367, 394* Richards, A. G., 75, 107*, 264,306*, 380,382,394* Richards, H. H., 54, 106* Richardson, C. D., 58, 109* Rick, J. T., 176, 180* Ricketts, J., 57, 58,61, 65, 66, 67,68, 69, 78, 79, 80, 81, 85, 87, 88, 90, 9 1 , 9 2 , 9 3 , 9 4 , 9 8 , 102* Riemann, J. G., 318, 324, 381, 394* Riess, R. W.,321, 332,334,338, 387* Rikmenspoel, R., 375,396* Rilling, G., 228,229,254* Rimon, A., 54, 108* Ringo, D. L., 341, 394* Ris, H., 329, 388* Ritchot, C., 57, 108* Robbins, W. G., 38 1, 394* Roberts, E., 171, 181 * Robertson, H. A., 49,49* Robertson, R. W.,58, 108* Robison, G . A . , 12, 15, 17, 31,35,36, 38,43*, 44*,47*, 48* Robison, W. G., Jr., 328, 353, 363, 370,371,380,394* Rock, G. C., 57, 108* Roeder, K. D., 258, 268, 282, 309*, 311* Rosati, F., 316, 317, 318, 319, 320, 322, 324, 325, 326,327,328,329,

Rosati, F.-cont. 330, 331, 333, 334, 335,336, 338, 339, 341, 342, 343,345,347, 348, 349, 350, 351, 352, 353,354, 355, 357, 358, 360, 361, 362, 363,364, 365, 366, 367, 369, 371, 377, 379, 380,381,385*, 386*, 394* Ross, A., 332, 394* Ross, J., 332,363, 370, 371, 394* Roth, L. E., 318, 394* Rothschild, Lord, 338, 381, 394*, 395 * Rothschild, H. A., 54, 108* Rottman, F., 65, 68, 87, 90, 107* Rowe, A. J., 341,343,390*, 394* Rowell, C. H. F., 260,280, 306* Royes, W. V., 57, 108* Rucker, W. B., 128, 179* Rudzisz, B., 67, 106*

S Sakai, A., 322,395* Salkeld, E. H., 234,238,250*, 254* Salpeter, M. M., 266, 307* Samli,,M. H., 37,47* Sances, A,, 304,307* Sandeman, D. C., 300,309* Sandlin, R., 176, 181* Sandoz, D., 326,395* Sang, J. H., 56, 104*, 108* Satir, P.,375,395* Sato, H., 332,391 * Sattelle, D. B., 259, 264, 267, 271, 273, 274, 276, 277,278,280,285, 286,288,296,309* Saxena, K. N., 194, 195, 196, 200, 202, 203, 206, 207,208,212,217, 247,254* Schalike, W.,332, 397* Schaller, G., 188, 210, 211, 212, 216, 217, 219, 220, 221,222,224,249, 255* Schoemaker, W.C., 38,44* Schogel, E., 3 , 4 9 * Scholes, J. H., 151, 164, 186* Scholz, H., 36,46*

AUTHOR INDEX

409

Smith, A. D., 34,48* Schorderet, M., 35,46* Schramm, M., 15, 16,37,42* Smith, D. S., 95, 96, 108*, 109*, 260, Schreiner, K., 375, 395* 264, 266, 271, 274,281,298, 299, 309*, 310*, 311* Schuberth, H. J., 94, 108* Schiirmann, F. W., 273,309* Smith, E., 61,65, 80,98, 105* Schwartz, I. L., 40,48* Smith, J. J. B., 188, 193, 196, 203, Scott, D. R., 223, 224,255* 206,208,238,251*, 255* Sedee, P. D. J. W., 56, 108" Smith, K. M., 185,203,255* Sellers, L. G., 68,70,87,88, 101* Smith, R. E., 38,44* Selmi, G., 322, 324' 325, 326, 327, Sogawa, K., 230, 231, 232, 233, 236, 240,241,255* 328, 329, 330, 331,335,336,342, 343, 345, 347, 348,349,350,351, Somlyo, A.P., 36, 38,44*,48* 352, 353, 354, 357,358, 360, 361, Somlyd, A. V., 36,38,44*, 48* 362, 363, 364, 365,366,367,369, Sorbo, B., 94, 108* 371, 377, 379, 380, 381, 385*, Spanner, S., 95, 101* Spencer, W. A., 151, 156, 157, 162, 386* 180* Sethi G. R., 75, 107* Sporn, M. B., 168, 179* Setsuda, T., 15,45* Springhetti, A:, 371,395* Sevhonkian, S., 224,254* Squarez, A., 72, 102* Seydoux, J., 38,44* Sribney, M., 54,109* Shanfeld, J., 36,47* Sridhara, S., 73,78, 109* Shapiro, B., 54, 108* Stamenovik, B. A., 34,45* Shapiro, D., 54, 106* Stauffer, J. F., 57, 101* Sharp, G. W. G., 40,45* Steele, J. E., 32,48*, 49,49* Sharp, P. L., 58,92, 104* Steinberg, D., 38,48* Shaw, J., 259,271,309* Stephens, R. E., 343,395* Shaw, J. E., 40,47* Sternlicht, E., 18, 37,45* Shaw, S., 151, 164, 180* Stevens, R. E., 341,395* Shay, J. W., 324,348,395* Stobbart, R. H.,259,274,309* Sheladci, M. L., 341,343,395* Streeto, J. M., 21,48* Shelley, R. M., 69,86,87,96, 108* Stride, G. O., 56, 109* Shigamatzu, H., 78, 105* Strong, F. E., 188, 204, 208, 209, Shigenaga, M., 322, 395* Shoger, R. L., 316,395* 214,220,222,255* Shoup, J. R., 331,395* Stryer, L., 176, 181* Siebold, C. T. von, 382, 395* Subrahmanyam, D., 88, 108* Siggins, G. R., 35,47* Sud, B. N., 334,395* Silvester, N. R., 375, 395*, 390* Sugioka, T., 334,335,396* Simmons, E. E., 348,395* Sundwall, A., 94, 108* Simpson, L. L., 34,47* Sutcliffe, D. W., 274, 3 1O* Singer, B., 40,44* Sutherland, E. W.,12, 15, 16, 17, 31, Singer, J. J., 34, 44*, 48* 34, 35, 36, 38, 39, 43*, 44*, 47*, Singer, T. P., 54, 108* 48 * Singh, K. R.P., 56,58, 108* Sylvester, F. S., 242, 255* Sjodin, R. A., 286,309* Skelton, C. L., 16,20,44*, 48 * T Slowiak, D., 188, 208, 215, 224,238, 239,244,253* Tahmisian, T. N., 324, 325, 334, Smallman, B. N., 63, 66,84,99, 108* 386*, 389* AIP--17

410

AUTHOR INDEX

Tahori, A. S., 58, 109* Tandler, B., 327,334, 395* Tasaki, I., 176, 181* Tates, A. D., 327, 351,388*,396* Taylor, E. W., 341, 343, 395* Taylor, J. H., 332, 396* Thiiry, J.-P., 341,384* Thomas, K. K., 75,76, 109* Thomas, M. B., 396* Thompson, R. F., 146, 147, 181* Thorn, N. A., 40,48* Thorson, B. J., 324,381,382, 394* Timms, A. R., 35,43* Tobias, J. M., 276, 310* Tomita, T., 35,42*, 43* Toppozada, A., 266,307* Tormey, J. M., 267,307* Treherne, J. E., 63, 95, 96, 107*, 108*, 109*, 257, 258, 259, 260, 261, 262, 264, 266,267,268,270, 271, 272, 273, 274,275,276,217, 278, 279, 280, 281, 282, 283, 284, 285, 286, 287, 288,289,290,291, 292, 293, 294, 296,298,299,300, 302, 303, 304, 308*, 309*, 310*, 311* Tsubo, I., 334,335,396* Tucker, L. E., 288, 293, 294, 295, 296,297,299,301,309*, 3 11 * Tunstall, J., 151, 164, 180* Twarog, B. M., 258, 268, 281, 282, 311*

U Uretz, R. B., 332, 391 * Usherwood, P. N. R., 259, 274, 31 1* Usinger, R. L., 215,251* Ussing, H. H., 285,286,307*, 31 1* Utiger, R. D., 37,48*

V Vale, W., 37, 48* van de Meene, J. G. C., 34,45*

Vanderzant, E. S., 57, 58,61, 109* Vane, J. R., 11,48* Van Herpen, G., 375,396* van Loon, L. C., 215,255* Vegni, M., 324,381,394* Venkitasubramian, T. A., 72, 78, 82, 105* Villeneuve, J.-L.,72, 78, 83, 101* Von Baumgarten, R., 164, 181* Vovis, G. F., 56, 61, 62, 65, 81, 99, 104*

W Wada, S. K., 326,396* Wahrman, J., 332,369,389* Walker, R. J., 63, 106*, 176, 180* Wang, C. M., 75,81, 109* Warner, F. D., 325, 336, 341, 343, 347,349,350,351,396* Watanabe, A., 176, 181* Wattal, B. L., 72,78,82, 105* Wearing, C. H., 207,255* Wechsler, W., 273, 309* Weidemann, H. L., 226, 239, 240, 255 * Weidler, D. J., 259, 262, 278, 279, 281,282,293,312* Weigt, W. A., 185, 186, 253* Weiss, B., 34, 48* Weiss, S. W., 54, 106* Wensler, R. J., 194, 255* Werner, G., 329, 334, 349, 365, 367, 370,396* Wetzel, R., 332, 397* Whelchell, K. E., 73, 78, 1l o * Whitaker, B. D. L., 34,45* Whitcomb, R. F., 220, 242, 251*, 255* Whitehead, A. T., 2,48* Whittaker, V. P., 96, 109* Wiersma, C. A. G., 147, 181* Wigglesworth, V. B., 32, 48*, 266, 268,298,305,3 12* Wilber, J. F., 37,48* Wilde, G. E., 208,241,25 1* Wilkes, A., 328,396*

41 1

AUTHOR INDEX

Williams, J. R., 199, 255* Williamson, J. R., 36,49* Willis, N. P., 67, 69, 85, 89, 90, 109* Wilson, E., 153, 155, 174, 181* Wilson, J. E., 179* Wilson, L., 341, 387* Winteringham, F. P. W., 68, 97, 99, 100, 109* WiSnieska, A., 74, 75, 78, 110* Wittenberg, J., 54, 109* Wlodawer, P., 74,75,78, 110* Wohlfarth-Bottermann, K. E., 226, 227,228,239,244,253*, 254* Wood, D. W., 276,312* Wozniak, A., 153, 155, 181* Wren, J. J., 72, 110* Wright, E. M., 285,307* Wroniszewska, A., 66, 105* Wullems, G. J., 32,45 * Wyatt, G. R., 49, 49*, 67, 70, 85,88, 104*, 1 lo*, 274,312*

Y Yamasaki, T., 258, 277, 278, 289, 312* Yasuzumi, F., 334, 335, 396* Yasuzumi, G., 322, 329, 330, 334, 335, 338, 348, 396* Young, J. A., 3 Young, 0. M., 54,106* Yund, M. A., 56, 61, 62, 65, 81, 99, 104* Yurkiewicz, W. J., 73,74, 78, 110*

Z Zelander, T., 341, 386* ZielMska, 2,M.,73,90, 110* Zirwer, D., 332, 397* Zylberberg, L., 325, 382, 397*


Subject Index A Abdominal ganglion, and learning, 153-5 Abdominal nerve cord, ion uptake, 95 Abdominal tissues, choline, 75 Acantholyda nemoralis, choline metabolism, 67, 73, 90 Accessory salivary gland, 236, 246 Acerentomon, sperm axoneme, 338 A . majus, 342 Acerentulus, sperm axoneme, 338 Acetylcholine and choline metabolism, vertebrates, 53-5 and choline synthesis, 91-100 and catecholamines, 34 extraneous application, 258, 282 metabolism, 63-6 Acetylcholinesterase, in choline metabolism, 84, 96 Acheta domesticus, choline metabolism in development, 57 lipids containing choline, 74, 75 phosphatidylcholine, 7 8 , 8 1 Acid phosphatase saliva, 210,215 sperm axoneme, 352 Acrosomal complex, sperm, 324-28, 382 Acrosternum hilare, and fungus, 241 Actin, sperm flagellum, 345, 375, 379 Action potentials and extra-axonal ions, 282-3,288-9 and Na+, 278 Active transport, in CNS, 300 Adelgidae, and phytopathogenicity, 22 1

Adelphocoris seticornis, salivary pectinase, 2 13 S-adenosylmethionine, in choline metabolism, 53-5 Adenyl cyclase, and cyclic AMP, 12, 18-19, 21, 29, 31-8,40,41 ADPase, sperm, 352 p -adrenergetic agents, and cyclic AMP, 35 Adrenocortieotropic hormone, and cyclic AMP, 36 Aedes aegypti, choline metabolism in development, 5 6 , 5 8 lipids containing choline, 1 2 phosphatidylcholine, 78 sphingomyelin, 8 3 substitutes, 59, 60 Aeneolamia, composition of saliva, 216 Afferent feedback, and learning, 164-6 Aggregation, slime mould, and cyclic AMP, 33-4 Agrotis orthogonia, choline in development, 57 Aiolopus strepens, sperm axoneme, 344,352 cell surface, 3 19,,323 Alanine, Hemiptera saliva, 218, 221 Aldosterone, mitochondria-rich cells, 40 Alkaline tetrazolium reaction, sulphydryl groups, 240 Altica ambiens alni, choline, 72 Amino acids, Hemiptera saliva origins of saliva, 236, 237 and phytopathogenicity, 21 8, 220-5 in sheath material, 206 in watery saliva, 2 16

413

414

SUBJECT INDEX

a-aminobutyric acid, aphid saliva, 2 18 Animals other than insects-cont. 7-aminobutyric acid, and salivary Dictyostelium discordeum, and gland stimulation, 6 cyclic AMP, 33-4 Amphibicorisae, feeding, 192 Diplopoda, sperm, 3 17 Amylamine, salivary gland stimulation, 6 Dog, learning, 143 Amylase Echinodermata, sperm, 341,343 and cyclic AMP, 37 Elasmobranchs, blood-brain saliva, 209, 214, 215 barrier, 302, 303 Fish, sperm 33 1 Anagasta, sperm capacitation, 38 1 Flagellates, flagellum, 349, 350 Anal cerci, and habituation, 15 1-2, Frog 156 body fluid composition, 275 Animals other than insects potential changes, 285,286 Acarina, sperm, 3 17 water permeability, 39 Amphibia Gastropoda blood-brain barrier,303 glycogen, 350 epithelia, permeability, 39 hyperpolarisation, 286 sperm, 349 paired sperm, 367 Amphioxus, sperm, 3 16 Hamster, blood-brain barrier, 302 Annelida Hirudo medicinalis, blood-brain blood-brain barrier, 258, 302, 303 barrier, 303 sperm, 3 16 Lamellibranch, blood-brain barrier, A plysia 303 learning, 162, 163, 164 Leech neurones, 277 blood-brain barrier, 303 Arachnida, sperm, 3 16, 3 17 glial cells, 273 Arthropods Loligo, choline, 94 and Reduviid saliva, 204 Mammal sperm, 316,327,331 blood-brain barrier, 300, 302, Branchiura, sperm, 3 16 3 03 Bufo marinus, cyclic AMP, 39-41 choline metabolism, 52, 59, 96 Bull, sperm, 374, 381 cyclic AMP, 12 Carcinas maenas, blood-brain learning and drugs, 169 barrier, 300, 303 saliva production, 3 Cephalopods sperm, 326,341,350,382 body fluid composition, 275 Marsupials, paired sperm, 367 excitation and conduction, 277. Merostomata, sperm, 31 6 Chilopoda, sperm, 3 16 Mollusca Chlamydomonas, flagellum, 347 heart, 5-HT, 11 Chordata, sperm, 3 16 sperm, 316 Cirrepedia, sperm, 3 16 Mouse Crab, shore, blood-brain barrier, blood-brain barrier, 302 3 00 choline metabolism, 94 Crustacea cyclic AMP, 18 blood-brain barrier, 258, 300, Myriapoda, sperm, 3 16, 3 17 302,303 Mystacocarida, sperm, 3 16 muscle fibre innervation, 147 Necturus, blood-brain barrier, 302 sperm, 316 Opilionids, sperm, 3 17 Decapoda, sperm, 330 Opossum, paired sperm, 369

SUBJECT INDEX

Animals other than insects-conl Pauropoda, sperm, 3 16 Protozoa flagellum, 341, 342, 343, 374, 375 learning, 176 Rat choline uptake, 95 learning, 114, 115, 126, 127-8, 168 stomach, 5-HT,11 Scorpion, sperm, 3 16 Sea urchin, sperm, 343, 345, 349, 3 74 Slime mould aggregation, and cyclic AMP, 33-4,41 Snake venom, 209 Squid, giant axon, 278, 291 Symphyla, sperm, 3 16 Toad, bladder, and cyclic AMP, 39-41 Teleost, blood-brain barrier, 303 Tetrahymena, cilia, 341, 343 Turritella terebra, paired sperm, 3 67 Vertebrates blood-brain barrier, 25 1,302,303 choline metabolism, 52, 53-5, 69, 71,82,89, 96 cilia, 350 cyclic AMP, 15 malignant trophoblastic cells, 304 and reduviid saliva, 204 salivary glands, 37 sperm, 316 Anoplura, sperm, 328, 369, 380 Antheraea pernyi, acetylcholine, 66 Antheraea polyphemus acetylcholine, 66 phosphorylcholine, 67 Anthonomus grandis, choline metabolism, 5 7 , 5 8 , 5 9 , 61 Anticholinesterases, 99-1 00 Anuraphis bakeri, choline metabolism, 73 Aphaniptera, sperm acrosomal complex, 324 axoneme, 338

415

Aphaniptera, sperm-cont. cell surface, 3 18 motility, 377,379 Aphididae, saliva composition, 209, 210,211 pectinase, 2 13, 2 14 Aphidoidea, saliva, 226-9,245 Aphids saliva composition, 211, 212,215, 216 excretion, 245 metabolites, 218-9,220, 221 methods, 185-7, 189 necrosis, 249 oxidases, 239,247 phenolic compounds, 249 phytopathogenicity, 2 17-25 stylet-sheath feeding, 194-202, 205,246 virus transmission, 242 sperm, 365 Aphid, black bean, feeding, 194 Aphis abbreviata, pectinase, saliva, 21 3 Aphis cardui, pectinase, saliva, 2 14 Aphis fabae, saliva feeding, 194, 195, 196 methods, 189 pectinase, 213 Aphis pomi, metabolites, saliva, 2 18-9 Aphis sambuci, metabolites, saliva, 2 18-9 Aphis sedi, pectinase, saliva, 213 Aphis spireacola, pectinase, saliva, 2 13 Aphis, woolly, saliva detoxicant function, 248 phytopathogenicity, 2 17 Aphoidea, feeding, 192 Aphrophora alni, salivary glands, 232 Aphrophora parallela, choline metabolism, 73 Apis mellifica, choline metabolism acetylcholine, 66 lipids containing choline, 75, 77 oxidation, 89 requirements, 92 Apterygota, sperm, 326, 327, 328, 329,338 Archips cerasivoranus, lipids containing choline, 73

416

SUBJECT INDEX

Arctia caia, choline metabolism, 66, 73,85 Arge pectoralis, lipids containing choline, 73 Arginine saliva, aphids, 2 18 sperm, 33 1,335 Argyrotaenia velutinana, choline in development, 57 Asopinae, feeding, 192 Asparagine, saliva, 2 16, 2 18 Aspartic acid, saliva, 216, 218, 221 Attagenus spp., choline in development, 56 ATP, conversion to cyclic AMP, 14 ATPase, snake venom, 204 sperm, 336, 343, 345, 346, 349, 363-7 371,375,377, 379 Atropos pulsatorium, sperm, 369, 370 Auchenorrhy ncha saliva composition, 209, 216 glands, 225, 233-4 pectinase, 213, 214 sperm, 365 Aulacaspis tetalensis, feeding, 198-9 Aulacorthum solani, saliva, 213, 249 Axoneme, sperm, 336-53, 374-80 axonemal matrix, 352-3 central sheath, 349 coarse fibres, 350-2 links heads, 349-50 microtubules, 338-49 and motility, 374-80

Betaine, and choline metabolism, 52, 53-5,59,63,89 Blaberus gigantea, neural fat body sheath, 280 Blatta orientalis, learning, isolated ganglion, 132, 158 Blattella germanica, choline metabolism, 57, 59, 61, 74, 89 Blattoidea, sperm, 324 Blood-brain barrier, 257-3 12 electrical aspects, nerves, 277-9 1 ionic basis, 277-8 neural fat body sheath, 278-81 neuronal function, 28 1-9 1 ionic composition, haemolymph and nervous tissues, 274-7 nervous tissue, organisation, 260-74 extraneuronal fat body deposits, 200-3 glial cells, neurones and extracellular spaces, 268-74 neural lamella, 264-6 perineurium, 266-8 radioactive ions and molecules, exchanges, 291-9 Bloodsuckers, saliva, 203, 204, 205, 206,215,238 BOL (Bromolysergic acid diethylamide), and salivary glands, 8-10 Bombyx mori choline metabolism in development, 5 7, 5 8 lipids, 73 phosphatidylcholine, 78, 85, 96 phosphorylcholine, 67 substitutes, 59, 61 haemolymph, 276 sperm, 34 1 Brain catecholamines and cyclic AMP, 34 choline, 75,77 see also Blood-brain barrier Bufotenine, and salivary gland stimulation, 7 Butterfly, sperm, 3 18

B B vitamins in development, 58 Bacillus rossius, sperm absence of mitochondria, 360 accessory flagellar bodies, 366,368 acrosomal complex, 327 axoneme, 345,346,352 cell surface, 323 centriole region, 335 motility, 377,368, 381 C nucleus, 330,331 Bacteria, and cyclic, AMP, 12 Caesium ions, and potential changes, Barium. and stimulation bv ADH. 40

SUBJECT INDEX

Calcium haemolymph, 275 -mediated action potentials, 278 see also Cyclic AMP Calligypona pellucida, saliva, 2 16, 224 Calliphora ery th rocep hala choline metabolism, 56, 72, 84 salivary glands, cyclic AMP, 1-49, 1-49 see Cyclic AMP Campodea, sperm, 325,342,344, 350, 361 Campodeidae, sperm, 347,349,351 Capsus ater, pectinase, saliva, 2 13 Carabids, sperm, 331 Carasius morosus extra-axonal sodium regulation, 3 02 extra neuronal potentials, 285, 288 fat body deposits, 260-3 ionic basis, electrical activity, 277, 278 ionic composition, nervous tissues, 275,276 neural fat body sheath, 278-8 1 neural lamella, 265-6 perineurium, 266,268 Carbohydrates saliva, 240-1 sperm, 352-3. 380 Cardiac cells, and cyclic AMP, 15 Cardophilus hemipterus, choline, 56 Carnitine, and choline metabolism, 52, 59-62, 70, 80,87, 99 Carnivores, saliva, 203-5,238 Catecholamines, and cyclic AMP, 34, 35-6,38 Cation exchange, blood and CNS, 258 CDP-choline (cytidinediphosphorylcholine), and choline metabolism, 53-5, 69, 86, 96, 97 Cecidogenesis, and saliva, 224-5, 249 Celerio euphorbiae, choline metabolism, 6 6 , 8 5 Cell membrane and choline, 52 and watery saliva, 201 Cell surface, sperm, 3 17-24 Cellulase, saliva, 197, 209 Centriolar region, sperm centriole, 332-3, 382

417

Centriolar region, sperm --con t. centriole adjunct, 329, 333-5, 365 initial segment, axoneme, 335-6 Ceramide, and choline metabolism, 5 3-5 Ceratatis capita choline metabolism, 7 1, 72 sperm, 337,344,346 Cerci, anal, and habituation, 15 1-2, 153-5 Cercopoidea, saliva, 214, 229, 232 Cercopidae, saliva, 2 16 Ceruraphis eriophori, saliva, 214 Ceruraphis viburnicola, saliva, 2 13 Chironomus spp., choline metabolism, 72 ChlGon dipteron, sperm, 325, 339 Chloride and diuretic hormone, 33 salivary glands, 3, 22, 24-5, 28,29, 31 Chlorogenic acid, aphid saliva, 219 Choline metabolism, 5 1-109 enzymes, 84-91 cholineacetylase and acetylcholinesterase, 84 choline, oxidation, 88-9 choline, synthesis, 89-91 phosphatidylcholine, hydrolysis, 8 7-8 phosphatidylcholine, synthesis, 85-7 lipid-soluble metabolites, 7 1-84 lysophosphatidylcholine, 82-3 phosphatidylcholine, 7 1-82 sphingomyelin, 83-4 metabolic role of choline, 91100 nutritional requirements, 5 5-63 in development, 55-8 substitutes, 59-63 vertebrates, 52, 53-5 water-soluble choline metabolites, 63-7 1 acetylcholine, 63-6 CDP-choline, 69 glycerylphosphorylcholine, 70-1 phosphorylcholine, 66-9 Choline ions, and potential changes, 283-7

418

SUBJECT INDEX

Chromatoid body, and centriole adjunct, 334 Chromosomes, sperm, 332 Chryptothnps latus, sperm axoneme, 351 Chrysomela crotchi, choline, 72 Chrysopa carnea, sperm axoneme, 339 Cicadellidae saliva, 213,230,233 sperm, 365 Cicadomorpha, saliva, 192, 233 Cicindela, sperm, 3 65 Cimex lectularius, salivary glands, 234, 235,238 Cimicidae feeding, 192 salivary glands, 235, 249 Cimicomorpha composition of saliva, 2 15 composition of sheath material, 206 feeding, 192, 193,203 salivary glands, 234, 238 Cinara, spp., pectinase, saliva, 213 Cixius nervosus, sperm, 364 Classical conditioning, 1 13, 162-4 Clitumnus, sperm, 360 Coccidae phytopathogenicity, 2 17 sperm, 332, 353, 363, 370, 374, 380,381 Coccoidea, sperm, 328 Cochliomyia hominivorax, choline, 56, 5 8 ,5 9 ,6 0 Cockroach abdominal nerve cord, 258 choline substitutes, 59 fat body deposits, 260 learning, isolated ganglia see Learning Cocoa capsid, phytopathogenicity, 217,220,223,225 Coleop tera choline metabolism analogues, 98 in development, 5 5-6 enzymes, 8 6 , 90 lipid-soluble metabolites, 71, 72, 7 8 ,8 2 ,83

Coleoptera-cont. choline metabolism-cont. phosphor ylch oline , 6 9 haemolymph, ionic composition, 275 sperm accessory flagellar bodies, 364 acrosomal complex, 324,328 axoneme, 337,348,350, 351 cell surface, 323 centriolar region, 336 mitochondria, 357,358,362 paired sperm, 367 Coleorrhy ncha feeding, 192 salivary glands, 225, 226, 231,233, 245 Collembola, sperm, 316, 324, 327, 338,354 Colour changes, mantids, 32 Conditioning, classical, 113, 162-4 Coreidae, pectinase, saliva, 2 14 Coreoidea, feeding, 192 Corixidae, feeding, 192 Cornus drummondi, (dogwood), and Acrosternum, 241 Corrodentia, sperm, 327 Cricket, house, sperm, 33 1,334 Cryptocerata, salivary glands, 234 Cryptomyzus nbis, saliva, 218-9 Ctenocephalus canis. sperm, 318, 376 CTPase, sperm, 352 Culex pipiens fatigans, choline metabolism, 71,72, 78, 82, 88 Culicid Diptera, sperm, 338 p-Cumaric acid, aphid saliva, 2 19 Cuticle choline, 75 formation, 239, 245 Cyclic AMP and Calcium, and hormone action, 1-49 Calliphora salivary glands, 2-5 5-HT, comparison 32-41 control of metabolism, 37-9 epinephrine and heart, 36 excitation-secretion coupling, 36-7 pre- and post-synaptic transmission, 34-6

SUBJECT INDEX

Cyclic AMP and Calcium-cont. SHT, comparison-cont. slime mould aggregation, 33-4 transporting epithelia, 39-41 5-HT-receptor interaction, 5-1 2 Intracellular messengers, 12-2 1 calcium, 19-2 1 cyclic AMP, 12-19 mode of action, 21-32 ion transport, 26-8 potential effect, 23-6 time course, 28-3 1 model of hormone action, 31-2 Cyclic nucleotides, and specificity of cyclic AMP, 16-18 Cycloheximide, and learning, 172-5 Cysteine, saliva, 21 1, 216,218 Cytidine pathway, 85 Cytochrome-c-oxidase, sperm, 35 7, 362,363,366

419

Diapheromera femorata, choline, 74 Dibutyryl derivative, cyclic AMP, 15-16, 34,39 Dictyoptera, ionic composition, haemolymph, 281 Diglyceride, and choline metabolism, 53-5 Dihydroxy phenylalanine, saliva, 223, 239,249 N-Dimethylaminoethanol, and choline metabolism, 59 Dimethylethylcholine, as choline substitute, 62 N-Dimethylglycine, and choline metabolism, 53-5 Dimethyliso-propylchohe, as choline substitute, 62 Dimethyltryptamine, and salivary gland,.7 Diplura, sperm, 316, 324, 327, 342, 344,347,349,350,354 Dipsocorinomorpha, feeding, 192 Diptera choline metabolism D analogues, 98 enzymes involved, 86 Dacus, sperm, 350,370 lipid soluble metabolites, 7 1, Dacus oleae, choline, 72 72-3,77, 78,82,83,84 Dactynotus, sp., saliva, 2 13 requirements, 55-8, 62, 92 Dahlbominus, sperm, 328 Dalbulus maidis, saliva, 2 13 synthesis, 90 Datana integerrima, choline, 73 water soluble metabolites, 69, DDD (dihydroxydinaphthyldi70 sperm sulphide), test for sulphydryl acrosomal complex, 327 groups, 240 axoneme, 337, 339, 341, 344, Delphacidae salivary glands, 232,233 346,348,350,351 mitochondria, 355,357,362 sperm, 365 Deltocephalidae, salivary glands, 230 non-flagellate sperm, 374 Deoxyribonucleoproteins, sperm Diuretic hormone nucleus, 332 and 5-HT, 11-12 Dermaptera, sperm, 336 Rhodnius, 33 Dermestes vulpinus, choline in DNA, sperm, 329,33 1,332,382 development, 56 Dolycoris baccarum, composition of Desheathing, and axonal response, saliva, 209,210, 215 279,286-7,288,289 Donnan equilibrium, and peripheral Detoxicant function, saliva, 246-7 diffusion barrier, 259,264 Development, choline in, 55-8 DOPA, saliva, 223,239,249 DFP (di-iso-propylphosphofluoridate) Dopamine, and cyclic AMP, 6,35 and acetylcholine, 99, 100 Dreyfusia spp., and galls, 220, 221

420

SUBJECT INDEX

Drosop h ila m elanogaster choline metabolism metabolic role, 92, 97, 99 oxidation, 89 lipid-soluble metabolites, 72, 8 1 requirements, 56, 58, 59,61, 62 substitutes, 64-5 sperm nucleus, 33 acrosomal complex, 327 axoneme, 338, 343, 344, 349, 350 centriole region, 333, 335, 336 mitochondria, 362 polymorphism, 382-3 spermatids, 370 Drosophila spp. choline in development, 57 sperm polymorphism, 383 Drugs, and learning, cockroach, 168-75 Dysdercus feeding, 194, 202-3, 207, 208 and fungus, 241 salivary composition, 2 12 Dysdercus koenigii, secretion of flange, 195 Dytiscus marginalis, paired sperm, 367,369

E Ecdysone, mode of action, 32 Ecdysterone, and cyclic AMP, 32 Efferent gating, and learning, 164-6 Egg, choline metabolism, 63-4, 72-4, 91, 92, 99 Electrical aspects, nervous function, 277-91 ionic basis, 277-8 neural fat body sheath, 278-8 1 neuronal function in experimental preparations, 28 1-9 intact systems, 281-9 Electron spin resonance, and membranes, 176 Embioptera, non-flagellate sperm, 373

Empoasca fabae saliva, 209, 210, 230, 231,232 Enzymes choline metabolism, 84-9 1 saliva, 37, 197, 203, 204, 206, 209-17, 237-9, 245 sperm axoneme, 352 Eosentomon, non-flagellate sperm, 3 74 Ephemeroptera, sperm, 327,338, 339, 354,363 Epinephrine, and cyclic AMP, 12, 21, 36, 37, 38 Epipharyngeal organ, and saliva, 2 12 EPSP, and learning, 157 Esterases, in saliva, 210, 212, 215 Ethanolamine as choline substitute, 59, 63 and salivary gland, 6 Ethylamine chain, and 5-HT-receptor interaction, 6-9 Ethylenediamine, and salivary gland, 6 Erannis tiliaria, choline, 73 Ericerus pela, choline, 73 Eriococcus, non-flagellate sperm, 370 Erisoma americanum, saliva, 213 Erisoma lanigerum, saliva, 213, 217, 218-9,220,221,248 Erythroneura limbata, salivary glands, 230 Euchistus, sperm, 359, 360 Eumecopus punciventris, saliva, 21 1, 24 1 Eurydema rugosa, stylet-sheath feeding, 195,207,208 Euphestia elutella, choline in development, 57 Euphestia kuehniella, choline in development, 57 Euxesta notata, choline, 72 Evolution of salivary function, Hemiptera, 207, 244-7 Excretion, salivary glands, 184,245

F Fat body choline, 75, 76 glycogen conversion, 32

SUBJECT INDEX

42 1

Ganglia- con t. isolated, learning, 11 1-81, see Learning Gelastocoris sp., sperm axonemes, 370 Genetics, sperm, 382-3 Geocorinae, feeding, 192 Geocorisae, feeding, 192,204 Giant axons cation gradients, 275 excitation and conduction, 277, 278 extraneuronal potentials, 282, 285, 288, 289-90 and glial cells, 272 Gibberellic acid, in saliva, 2 16 Glands and ducts, salivary, Hemiptera, 225-35 Aphidoidea, 226-9 Fulguromorpha, 232-3 Heteroptera, 234-5 Jassomorpha, 229-32 other Auchenorrhyncha, 233-4 Glial cells choline uptake, 95 organisation, 268-74 and Sodium regulation, 279, 300, 304-5 Glossina morsitans, choline, 71, 73 Glucagon, and cyclic AMP, 12, 38 Gluconeogenesis, renal, and cyclic AMP, 39 Glucose, and cyclic AMP, 38 Glutamic acid, Hemipteran saliva, 216, 218,221 Glutarnine, Hemipteran saliva, 216, 218,221, Glyceraldehyde-3-phosphate dehydrogenase, sperm axoneme, 352 Glycerylphosphorylcholine, metabolism, 53-5, 70-1 Glycine G aphid saliva, 2 18 Galleria mellonella and choline metabolism, 53-5 choline metabolism, 66, 67, 68, 74, Glycogen 75, 78,82 deposition, nervous system, 305 neural lamella, 264 sperm axoneme, 346,348,350 Galls, and Hemipteran saliva, 184, to trehalose, hormones, 32 191,211,217, 220-5 Glycogenolysis, and epinephrine, 36 Ganglia Glycolysis, anaerobic, sperm, 359, K' depolarisation, 28 1 363,380

Fat cells, and dibutyryl cyclic AMP, 16 Fat body sheath, neural, role, 278-81 Fat body deposits, extraneural, 260-3 Fatty acids, free, and cyclic AMP, 38 Feedback cyclic AMP and Calcium, 19-21,36, 40,41 and learning, 132, 164-6 Ferrulic acid, aphid saliva, 2 19 Flagellar apparatus, sperm, 367-74 non-flagellate sperm, 370-4 paired sperm, 367-9 two axonemes, 369-70 Flagellar bodies, accessory ordered, 363-7 Flagellar filament, axial, sperm, 336-53 Flagellar motion, sperm, 335, 336, 374-82 Flea, sperm, 318 Flight muscle, choline metabolism, 76, 77,83 Fluid secretion epithelia compared with penneurium, 267 regulation, 3 7 Fluorescent dye techniques, and membranes, 176 Formica rufa, oxidation of choline, 89 Fulguroidea, pectinase, saliva, 2 14 Fulguromorpha, saliva, 192, 232-3, 236,240,246 Fungal disease, and saliva, 241-2 Fructose-diphosphate aldolase, sperm axoneme, 352 Fruit-fly, sperm, 3 18

422

SUBJECT INDEX

Glycolytic pathway, sperm axoneme, 352 Glycoproteins, sperm, 381 Gold complex, sperm, 322, 324, 325, 365 Granular cells, and Sodium transport, 40 Grape phylloxera, saliva, 2 1 6 , 2 17 Gryllotalpa gryllotalpa, sperm, 343, 345,376,379 Gryllus, sperm axoneme, 342, 345 Gryllus bimaculatus, choline, 1 4 GTPase, sperm, 352 Gustatory sensilla, and saliva, 2 12 Gut, choline, 7 5 ,76

H Habituation, electrophysiology, 150-7 Haematopinus suis, sperm, 333, 369, 376 Haemocytes, and extraneural space, 262 Haemolymph choline, 75 ionic composition, 274-7 Harmala alkaloids, and salivary gland stimulation, 8-9 Harmaline, and salivary gland stimulation, 8 Heart, and cyclic AMP, 16, 21,32, 41 Heliothis zea, choline metabolism in development, 57 enzymes, 8 5 ,8 9, 90 lipids, 74 water-soluble metabolites, 67, 69 Hemiodoecus veitchi, salivary glands, 226 Hemiptera choline metabolism, 56, 58,71,73, 78,82 saliva see Saliva sperm, 327,355,381 Heteroptera saliva composition, 205, 207, 208, 209,213,214,216

Heteroptera-con t. saliva-cont. evolution, 245-1 glands and ducts, 184, 225, 234-5 feeding, 191, 192, 193-6 methods, 189 origin, 236,241 sperm, 327,370 Histamine, and salivary gland, 6 , 7 Histidine saliva, 218,221 sperm, 33 1 Histones, sperm, 331, 335 Homocysteine, and choline metabolism, 52, 53-5 Homoptera saliva composition, 205,206,207,212 evolution, 245-7 feeding, 191, 192, 194-6 glands and ducts, 184-5,225,234 origins, 236, 237-8 sperm, 327,345,365,370 Honey bee, sperm, 324,338 Hormone action, role of cyclic AMP and Calcium 1-49, see Cyclic AMP Hormone, plant, in saliva, 216 Housefly, choline metabolism, 62, 63, 69, 70, 82 5-HT Calliphora salivary glands, 2-5 compared with other hormones, 32-41 control of metabolism, 37-9 epinephrine and heart, 36 excitation-secretion coupling, 3 6-7 pre- and post-synaptic transmission, 34-6 slime mould aggregation, 33-4 transporting epithelia, 39-41 intracellula messengers, 12-2 1 calcium, 19-2 1 cyclic AMP, 12-19 mode of action, cyclic AMP and Calcium, 21-32 effect on potential, 23-6

423

SUBJECT INDEX

5-HT-con t. Inotropism, and cyclic AMP, 36 mode of action-cont. Insecticides and blood-brain barrier, 259 ion transport, 26-8 and choline metabolism, 52, 99, time course, 28-31 100 model of hormone action, 3 1-2 Instrumental learning, 113-5, 157-62, receptor interaction, 5-12 164 Hyalophora cecropia, choline metaIntestine, and cyclic AMP, 35 bolism Intracellular messengers enzymes, 85, 88 calcium, 19-2 1 lipid-soluble metabolites, 75, 76 cyclic AMP, 12-19 water-soluble metabolites, 66, 67, Inulin penetration, nervous system, 70 268 Hyalopterus pruni, metabolites, saliva, Ion transport 2 18-9 abdominal nerve cord, 95 Hyaluronidase, saliva, 204,210 and cyclic AMP, 38, 4 1 Hydrocorisae, saliva, 204, 192, 23 1, and diuretic hormone, 33 235 intact and headless preparations, Hydrolysis of phosphatidylcholine, 118-28 87-8 5-hydroxytryptamine, see 5-HT isolated ganglion, 128-136 5-hydroxytryptophane, and salivary other CNS lesions, 136-7 dand stimulation. 7 and electrical activity, nerve cells, Hylemya antiqua, cholhe metabolism, 277-8 57.73 salivary gland, 22-5,26-8, 3 1 Hyalobius pales, choline metabolism IPSP’s, and learning, 157, 162 enzymes, 8 5 , 8 9 , 9 0 Isethionate, and 5-HT, salivary glands, Homoptera 67, 69 28,29 Hymen opt era Isoptera, sperm, 328,329 choline metabolism lipid-soluble metabolites, 71, 73, 82 phosphoryl choline, 69 J requirements, 92 synthesis, 91 327,345,365,370 Japyx, sperm axoneme, 350 Hyperin, aphid saliva, 2 19 Japigidae, sperm axoneme, 338 Hyphantria cunea, lipids containing Jassoidae, saliva, 209, 210 choline, 74 Jassoidea, saliva, 214, 229-30, 245 Jassomorpha, saliva, 192,229-32, 233, 242 Jopeicidae, saliva, 2 15 I Juvenile hormone, mode of action, 32

Imaginal discs, choline, 75 Indole acetic acid, saliva, and phytopathogenicity, 219, 220, 222, 223-5,249 p-indolyl acetic acid (IAA), in saliva, 2 16-7 Inhibition, central, and learning, 162

K Kalotermes, sperm, 363 K . flavicollis, 371, 373

424

SUBJECT INDEX

Kalotermitidae, non-flagellate sperm, 37 1 Kermes, sp., non-flagellate sperm, 370 Kidney, adenyl cyclase, 21, 38 Krebs cycle, sperm, mitochondria, 359 L Lacerateand-flush feeding, Hemiptera, 191-3, 202-3, 207, 208, ,217, 220,222,246 Lactic acid, sperm axoneme, 353 Lactic dehydrogenase, sperm axoneme, 352 Laodelphax striatellus, salivary glands, 233 Larva, choline requirements, 92 Lasioderma serricorne, choline requirements, 5 5 ,5 6, 60 Leaf hoppers, saliva, 217, 240 Learning and memory, isolated ganglia, 11 1-8 1 behavioural investigations, 1 18-48 ganglionless P and R preparations, 137-40 intact and headless preparations, 118-28 isolated ganglion, 128-136 other CNS lesions, 136-7 P and R behaviour, ganglionic innervation of legs, 140-6 P and R behaviour, ganglionless legs, 146-8 concept of learning, 113-5 electrophysiological studies, 150-67 classical conditioning, 162-4 habituation, 150-7 instrumental learning, 157-62 newer approaches, 164-7 histological and anatomical, cockroach, 149-50 ganglion transplantation, 150 metathoracic ganglion, mapping, 149-50 “model” systems, use, 1 17 molecular approaches, 167-76 drugs, cockroach, 168-75 speculations, 175-6 reformation of concept, 1 15-7

Lepidoptera blood-brain barrier haemolymph, ionic composition, 275 neural lamella, 264 choline metabolism in development, 55-6 enzymes, 8 6 , 9 0 lipid-soluble metabolites, 7 1,73, 78,83 metabolic role, 92 wat er-soh ble metabolites, 6 6, 68, 69, 77 sperm cells acrosomal complex, 327 axoneme, 342,347,348,35 1 capacitation, 38 1 cell surface, 318, 320,322 mitochondria, 355 nucleus, 331 spermatogenesis, 382 Lepisma saccharina, lipids containing choline, 74 Lepismatidae, sperm cells, 349, 35 1, 367,369 Leucine, aphid saliva, 21 8 Liocoris lineolaris, pectinase, saliva, 214 Lipases and cyclic AMP, 38 saliva, 2 15 Lipids containing choline, 72-6 lipid-soluble choline metabolites, 7 1-84 lysophosphatidylcholine, 82-3 phosphatidylcholine, 7 1-82 sphingomyelin, 83-4 in saliva, 240-1 Lithium ions, and potential changes, 283-6 Liver, and cyclic AMP, 12, 38, 41 Locust classical conditioning, 164 fat body deposits, 260 learning eye, 151, 152 leg position, 118-9, 122, 141, 157-9

SUBJECT INDEX

Locust-cont. sperm cells capacitation, 381 cell surface, 3 18, 324 Locusta migratoria blood-brain barrier glial system, 273 haemolymph, 276 choline metabolism in development, 57 lipids containing choline, 74 phosphatidylcholine, 85, 96 Lucilia cuprina, lipids containing choline, 73, 76 Luteinking hormone, and cyclic AMP, 37 Lygaidae, saliva composition, 205, 209, 210, 211, 214 feeding, 191,196, 202,203 galls, 224 glands, 235, 237 methods, 188 Lygaeoidea, feeding, 192 Lygus spp., saliva, 224,241 L . disponsi, 209, 210, 214, 215, 238 L. elisus, 223 L. hesperus, 209, 212, 213, 222-3 L. pratensis, 213, 235 Lysine Hemiptera saliva, 2 18,22 1 sperm cells, 326,331 Lysophosphatidylcholine, metabolism, 53-5,72-6,82-3,84

M Machilidae, sperm cells, 349,354 Machilis, sperm cells, 330 M. distincta, 364 Macrosiphoniella millefolii, pectinase, saliva, 2 13 Macrosteles fascifrons, salivary transmission of disease, 242-3,244 Magnesium and ADH, 4 0 and blood-brain barrier, 259

425

Magnesium-con t. and sperm malformation, 383 Malacosoma americanum, acetylcholine, 66 Malathion, and acetylcholine, 99, 100 Malpighian tubules choline, 75 diuretic hormone. 33 Mallophaga, sperm cells, 327, 329, 351.369 Manduca sex ta blood-brain barrier electrical aspects, 278,280, 285, 288.302 haemolymph, ionic composition, 276 nervous tissue, organisation, 264,266-7,273 lipids containing choline, 76 Manganous ions, and action potential, 278 Mapping, metathoracic ganglion, 149-50 Mastotermitidae, non-flagellate sperm, 371 Matsucoccus bisetosus, non-flagellate sperm, 370 Mecoptera, sperm cells acrosomal complex, 324 axoneme, 338,339,341 mitochondria, 355 nucleus, 328 Megoura viciae accessory flagellar bodies, sperm, 364 metabolites, saliva, 21 8-9 Melaphis rhois, pectinase, saliva, 214 Membranes conformational changes, and learning, 176 sperm cells, 322 Membrane permeability, and cyclic AMP, 38 Membrane potential, and Na+ concentration, 289 Memory and learning, isolated ganglia, 1 11-8 1, see Learning Menopon gallinae, sperm axoneme, 35 1

426

SUBJECT INDEX

Mercaptoethanol, induction of biflagellate sperm, 369 Mesothoracic, ganglion, and learning, 124-5 Mesothoracic leg, and learning, 1 19, 144 Messengers, intracellular calcium, 19-21 cyclic AMP, 12-19 Metabolism, control, and cyclic AMP, 37-9 Metathoracic ganglion and learning, 124-5, 132, 150, 158-62, 177 mapping, 149-50 Metathoracic leg, and learning, 122, 124, 132, 135,144,157 Methionine aphid saliva, 2 18 choline metabolism, 52,53-5,59 N-methylated ethanolamines, as choline substitutes, 63 LY and p-methylcholines, as choline substitutes, 62 Milkweed bug, saliva, 188 Miniature end-plate potentials, and cyclic AMP, 34-5 Miridae, saliva composition, 209, 210,213, 215 feeding, 192, 193,203,207,208 glands, 235 and phytopathogenicity, 2 17, 220, 222 Miris dolabratus, composition of saliva, 213,215 Mitochondria -rich cells, and Na* transport, 4 0 sperm cells absence, 360-3 accessory flagellar bodies, 363-7 axoneme, 353,363 derivatives, 354-60 normal mitochondria, 354 polymorphism, 382 Model of hormone action, and cyclic AMP, 3 1-2 Molecular approaches, learning, 167-76 drugs, cockroach, 168-75 speculations, 175-6

Monoctenus juniperinus, lipids containing choline, 73 N-Monomethylaminoethanol, and choline metabolism, 59 Monomolecular films, calcium, displacement by ADH, 40 Moths, sperm cells, 318, 324,381 Motility, spermatozoa, 345, 352, 367, 370,371 capacitation, 38 1-2 mechanisms, 374-80 metabolism, 380-1 Motor output, and learning, 164-6 Motor neurones, release of ACh, 34 Murgantia histrionica, sperm mitochondria, 356 Musca domestica, choline metabolism enzymes involved, 84, 85, 86-8,90, 91 lipid-soluble metabolites, 73, 75, 76,77, 78, 79,80,82, 83 metabolic role, 92, 93-5, 96,97,98, 99 nutritional requirements, 57, 58, 59,61 water-soluble metabolites, 63, 64-5, 66,67,69 Muscle choline, 75,76 heart and visceral, regulation, 32 smooth, and cyclic AMP, 35, 41 Mycetophilid Diptera, sperm axoneme, 338 Mycoplasmal particles, Hemipteran saliva, 242-3,250 Myosin, in flagellum, 375 Mystacides azurea, sperm axoneme, 347 Myzus ascolonicus, saliva, 190, 218-9 Myzus cerasi, Myzus cerasi, saliva, 214, 218-9, 221 Myzus persicae choline in development, 57,58 saliva, 200,213,227,239,240,244

N Naringerin, aphid saliva, 219 Naucoris cimicoides, saliva, 205

427

SUBJECT INDEX

Nematospora coryli, and Acrosternum, 24 1 Nematospora jossypii, and Dysdercus, 24 1 Nemoura cinerea, sperm axoneme, 346 Neodiprion sertifer, lipids containing choline, 73 Nepa, sperm, 370 Nephotettix cinticeps, saliva, 230, 240 Nephrotoma, sperm axoneme, 345 Nephrotoma sodalis, lipids containing choline, 73 Neural fat body sheath, role, 278-81 Neural lamella as ion barrier, 95 organisation, 264-6,273 Neuronia. sperm axoneme, 339 Neurones and glial cells, organisation, 268-74 neuronal function experimental preparations, 289-9 1 intact nervous systems, 28 1-9 Neuroptera, sperm cells, 328, 329, 336,339,341,348,35 1 Neurotoxin, phospholipase as, 210 Neurotransmitter substances, and choline, 53,98 Noradrenaline, and cyclic AMP, 35 Norepinephrine, and K+ efflux, 38 N o tonec ta glau ca salivary glands, 23 1,235 sperm axoneme, 339 Nucleotides, cyclic, and specificity of cyclic AMP, 16-18 Nucleus, sperm, 328-32 chemical characteristics, 33 1 physical characteristics, 33 1-2 shape, 328-29 submicroscopic structure, 329-3 1

0 Oligosaccharases, saliva, 209 Oncopeltus fasciatus choline metabolism, 63, 66, 73, 78 saliva composition, 209, 2 10, 2 1 1, 2 12

Oncopeltus fasciatus-cont. saliva-con t. enzymes, 238 feeding, 194, 202-3,204,205 glands, 234,235 Operant learning, 113-5, 157-62, 164 Orthop tera choline metabolism enzymes, 86 lipid-soluble metabolites, 7 1, 74, 78, 82, 83 requirements, 55-6, 92 water-soluble metabolies, 69, 70 haemolymph, ionic composition, 28 1 sperm cells acrosomal complex, 324, 326 axoneme, 348,350,352 cell surface, 323 centriolar region, 336 genetics, 383 nucleus, 33 1, 332 Oryctes, sperm, 328 Osmotic pressure, and sperm motility, 381 Ostrinia nubilalis, acetylcholine, 66 Oxidases, Hemipteran saliva, 220, 238-9,246,247 Oxidation of choline, 88-9 Oxygen, and sperm, 380 Oxytocin, and cyclic AMP, 39

P Paleacrita vernata, lipids containing choline, 74 Palmitoleic acid, and choline metabolism, 77 Palorus ratzeburg’, choline requirements, 56,59,60 Pancreas, regulation by secretin, 37 Pancreozymin, stimulation of pancreas Panorpa annexa, sperm axoneme, 339 Parafilms, use in Hemipteran feeding, 185, 189,194, 197,200,212 Parasympathetic stimulation, salivary glands, 37 Parathion, and acetylcholine, 99

428

SUBJECT INDEX

Parathyroid hormone, and cyclic AMP, 14,21, 38 Parlatoria oleae, sperm, 353,370 Parotid glands and calcium 37 and dibutyryl cyclic AMP, 16 Pathogens, and saliva, 241-4 Pavlovian conditioning, 1 13, 162-4 Pectinase, in saliva, 197, 2 13 Pectinophor gossypiella, choline requirements, 57 Pectinpolygalacturonase, in saliva, 209,212,215,220,222 Pediculus, sperm, 369 Peloridiidae, salivary glands, 233, 245, 246 Pentatoma rufipes, saliva, 205, 235 Pentatomidae, saliva, composition, 205, 208, 209, 210, 21 1,214,215,216 feeding, 202, 203 glands, 234,235,237 Pentatomoidea, feeding, 192 Pentatomorpha, saliva, composition, 205, 206, 207, 208, 210,211 feeding, 191, 192, 193, 195, 196, 203 glands, 234,235,246 oxidases, 239, 247 PAS reaction, 241 Performic acid-alcian blue, for sulphydryl groups, 240 Perineurium, and blood-brain barrier, 264-5, 266-8, 273, 285, 290, 29 1, 300-5 Perineurial cleft, “tight junctions”, 95 Periodic acid-Schiff (PAS) reaction, Hemipteran saliva, 240, 241 Periphyllus negundinis, pectinase, saliva, 2 13 Periplaneta arnericana blood-brain barrier cation exchange, blood and CNS, 25 8 extra-axonal sodium regulation, 302,304 extraneuronal potentials, 282-3, 285.286.288.289.290 , _ , ,

Periplaneta americana-cont. blood-brain barrier-con t. fat body deposits, 260, 281 glial system, 268-74 glycogen deposition, nervous system, 305 ionic basis, electrical activity, 277,278 ionic composition, haemalymph, 275 ionic composition, nervous tissues, 275, 276 nervous tissues, exchange properties, 301 neural lamella, 264,266 perineurium, 266-9 trehalose and glucose uptake, 258 choline metabolism acetylcholine, 63, 95 acetylcholinesterase, 96 glycerylphosphorylcholine, 70 lipids containing choline, 74 phosphatidylcholine, 85-8 phosphorylcholine, 67 5-HT, 2 and Hemipteran saliva, 204, 205 learning and memory, isolated ganglia, 111-8 1 see Learning sperm, motility, 380, 38 1 Peroxidase penetration, nervous system, 262, 266, 267, 273, 279, 285, 302 in saliva, 215, 238-9, 244,245,247 PH of saliva, 217 and sperm motility, 380 Phagostimulants, and host specificity, 248 Phasmida, ionic composition, haemolymph, 275 Phasmoidea, sperm cells absence of mitochondria, 360, 363 accessory flagellar bodies, 365 acrosomal complex, 327 axoneme, 342, 348, 350, 351, 352 cell surface, 323 centriolar region, 336 Phenols, Hemipteran saliva, 223, 224

SUBJECT INDEX

Phenol/phenolase system, Hemipteran saliva, 247, 249 Phenolases, Hemipteran saliva, 223, 224 Phenolase/peroxidase system, Hemipteran saliva, 2 10 Phenolic compounds, Hemipteran saliva, 219, 221, 247, 248,249 Phenylalanine, Hemipteran saliva, 2 16, 223,224 Phenylalanine/Tyrosine/DOPA, , Hemipteran saliva, 2 11 Phenethylamine, salivary gland stimulation, 7 Philaenus, sperm, nucleus, 331 Philaenus spumarius, salivary glands, 232 Phloridzin, aphid saliva, 219 Phormia regina, choline metabolism enzymes, 85-91 lipid-soluble metabolites, 73, 78, 80,81,84, metabolic role, 96, 97,98 nutritional requirements, 57, 58, 59,61,62 water-soluble metabolites, 64-5, 67, 69,70 Phorodon humili, pectinase, saliva, 213 Phosphatidylcarnitine, metabolism, 87 Phosphatidylcholine, and choline metabolism, 7 1-82 enzymic synthesis, 85-7 hydrolysis, 87-8 and metabolic role of choline, 92, 94, 95, 96,98, 100 in vertebrates, 5 3-5 Phosphatidyl DMAE, and choline metabolism, 53-5 P h o s p ha t id y l e t h a n olamine, and choline metabolism, 53-5,77, 79, 96 Phosphatidyl-MMAE, and choline metabolism, 53-5 Phosphatidylserine, and choline metabolism, 53-5 Phosphodiesterase, and cyclic AMP, 12-18,27, 35 Phosphofructokinase, sperm axoneme, 352

429

Phospholipase, saliva, 204, 210 Phospholipids and choline metabolism, 52-3 in saliva, 240 Phosphorylase and cyclic AMP, 38 and epinephrine, 36 in saliva, 210,215 sperm axoneme, 352 Phosphorylase b kinase, and cyclic AMP, 17 Phosphorylcholine, and choline metabolism, 53-5, 66-9, 92, 97 Phylloxera, galls, 22 1, 222, 224 Phylloxeridae, salivary glands, 228-9, 245 Phytopathogenicity, and Hemipteran saliva, 2 17-25 Phytophaga rigidae, lipids containing choline, 71,73 Phytophagous insects blood-brain barrier, 280, 288, 302 saliva, 184, 192, 193, 196, 197, 203, 204, 207, 210, 212, 224 Pikonema alaskensis, lipids containing choline, 73 Pilocarpine, and salivation, 188 Pissodes strobi, lipids containing choline, 72 Plasma membrane, labilisation, and cyclic AMP, 4 1 Platymerus rhadarnanthus, saliva, 204, 205,210,238 Plecoptera, sperm, 327,341,365 Plodia interpunctella, choline metabolism, 74,78 Poeciloscytus unifasciatus, pectinase, saliva, 2 13 Polymorphism, sperm, 382-3 Polyphenol oxidase (PPO), Hemipteran saliva, 215, 223, 238, 240, 245 Polysaccharide, sperm, 348, 352, 359, 365,380 Potassium and blood-brain barrier, 259, 272, 274 and cyclic AMP, 38 and extraneuronal potentials, 281-91

430

SUBJECT INDEX

Potassium-con t. haemolymph, 275 and osmotic gradients, salivary glands, 22,24 and resting potential, 277 in saliva, 3 and sperm malformation, 383 uptake, abdominal nerve cord, 95 Potentials extraneuronal, 282-9 membrane, and cyclic AMP, 38 miniature end-plate, and cyclic AMP, 34-5 salivary glands, effect of 5-HT and Cyclic AMP, 23-6,27 Predatory insects, saliva, 193, 203-5, 208,210,215 Procaine, and action potential, 278 Prociphilus tessellatus lipids containing choline, 73 pectinase, saliva, 214 Prostaglandins, and Calsium, 40 Protamines, sperm nucleus, 333 Protein synthesis increased after learning, 168 inhibition by cycloheximide, 174 Proteinases, in Hemipteran saliva, 204, 210,215,219 Protenor, sperm genetics, 383 Prothoracic ganglion, and learning, 124-5, 129, 132, 136, 137, 138-40, 149, 169, 172-3 Prothoracic leg, and learning, 124, 125, 129, 135, 138-40, 144 Protocatechuic acid, in aphid saliva, 219 Protoparce sex fa. choline metabolism, 67,68,69,74, 82 Protura, sperm acrosomal complex, 327 axoneme, 338,342,343 non-flagellate sperm, 374 Pseudococcus obscurus, non-flagellate sperm, 370 Pseudosarcophaga affinis, choline in development, 57 Psocidae, sperm axoneme, 338 Psocoptera, sperm, two axonemes, 369 Psychodidae, sperm, 327, 355, 374

Psyllidae saliva, 2 14, 2 17 sperm nucleus, 33 1 Pterocomma spp., pectinase saliva, 213 Pterygota, sperm acrosomal complex, 324 axoneme, 349 mitochondria, 354,355,363 Ptinus tectus, choline in development, 56 Purkinje cells, cerebellar, and cyclic AMP, 35 Puto, non-flagellate sperm, 370 P. albicans, 370 Pyrausta nubalis, choline in development, 57 Pyrrhocoridae, saliva composition, 21 0 feeding, 191, 196,202,203 principal gland, 237 Pyrrhocoris, sperm, two axonemes, 370 Pyrrhocoris apterus, saliva, 210, 215, 244

Q Quaternary ammonium compounds, uptake by abdominal nerve cord, 95 Quercitin, aphid saliva, 2 19 Quercitrin, aphid saliva, 219 Quinones, Hemipteran saliva, 222, 223,247

R Radioisotopes, and salivation, 189, 190 Receptor-5-HT interaction, 5-1 2 Reduviidae, saliva composition, 2 10 feeding, 192,203,204,208 glands, 235

SUBJECT INDEX

431

Reduviidae, saliva-cont. Rutin, aphid saliva, 219 and rickettsia1 diseases, 250 Reduvioidea, feeding, 193 Reduvius personatus, saliva, 205 Renal tubules, and cyclic AMP, 14 S Reproductive organs, acetylcholine content, 66 Saldidae, feeding, 192 Resting potential, and extra-axonal Saliva, Hemiptera, 183-255 ions, 277, 288 composition and function, 205-1 7 Reticulitermes, sperm, 354 sheath material, 205-8 R. lucifugus, 371 373 watery saliva, 208-1 7 Rhabdophaga swainei, lipids conevolution, 244-7 taining choline, 7 1, 73 feeding by carnivores, 203-5 Rhopalosiphum rhois, pectinase, lacerate-and-flush feeding, 202-3 saliva, 2 13 methods, 185-90 Rhincoris carmelita, saliva, 205 modes of feeding, 190-3 Rhinotermitidae, non-flagellate sperm, origins, 236-41 371 accessory gland, 236 Rhodnius prolixus principal gland, 237-8 diuretic hormone, 33 salivary carbohydrate and lipid, 5-HT and Malphighian tubules, 240-1 11-12 sources of oxidases, 238-9 saliva sources in Homoptera, 239-40 composition, 205,206, 208 phytopathogenicity, 2 17-25 feeding, 188, 193, 192, 196 salivary glands and ducts, 225-35 glands, 234,235 Aphidoidea, 226-9 origins, 238,241 Fulguromorpha, 232-3 Rhynchota, sperm Heteroptera, 234-5 accessory flagellar bodies, 364 Jassomorpha, 229-32 axoneme, 339,342 other Auchenorrhyncha, 2 3 3 4 cell surface, 328 stylet-sheath feeding, 194-202 mitochondria, 356,359,360 as vehicle for pathogens, 241-4 non-flagellate sperm, 370 Salivary glands, cyclic AMP and two axonemes, 369,370 Calcium, 1-49, see Cyclic AMP Rhynchotoidea, sperm, 35 1, 269-70, Saldidae, feeding, 192 380 Sappaphis, mali, metabolites, saliva, Ribonucleoproteins, in sperm centriole 218-9 adjunct, 329 Rickettsial diseases, Sarcophaga, sperm axoneme, 353 transmission, 250 Sarcophaga bullata, choline metaRNA bolism, 75,85 increased, after learning, 168 Sarcoplasmic reticulum, permeability, sperm nucleus, 33 1 and cyclic AMP, 36 Robinetinaglycone, aphid saliva, 2 19 Sarcosine, and choline metabolism, Romalea microptera, ionic composi53-5 tion, nervous tissues, 275,276 Sarcosomes, flight muscle, choline Royal jelly, acetylcholine content, 66, metabolism, 76 92 Scale insects, sperm nucleus, 330 Rubidium ions, and potential changes, Schistocerca gregaria 283-6 choline metabolism, 57, 75

432

SUBJECT INDEX

Schistocerca gregaria-cont. fat body deposits, 260 learning, leg position, 157-9 Schizolachnis pini-radiata lipids containing choline, 73 pectinase, saliva, 2 13 Sciara coprophila, sperm acrosomal complex, 327 axoneme, 338,340,343 capacitation, 38 1 centriole, 334 flagellum, 367 Scratch-and-suck feeding, 19 1, 2 17 Secretin, and fluid transport, pancreas, 37 Sensilla, gustatory, and saliva, 2 12 Sensory input, and learning, 164-6 Serine and choline metabolism, 5 2 , 5 3 4 Hemipteran saliva, 2 18,22 1 Serotin, snake venom, 204 Shock-avoidance learning, cockroach, 169-7 1 Simple problem solving, 1 13-5, 157-62, 164 Sitodrepa panicea, choline in develop ment, 5 5 Smooth muscle, and cyclic AMP, 35, 41 Sodium and cyclic AMP, 36,39-40 and fat body cells, 262 in haemolymph, 275 and neural fat-body sheath, 278-8 1 and neuronal function, 282-9, 299, 300, 302-4 and peripheral diffusion barrier, 259 and resting potential, 277 uptake, abdominal nerve cord, 95 Sodium pentobarbitol, and learning, 169 Sperm cells, 3 15-97 accessory ordered flagellar bodies, 363-7 acrosomal complex, 324-28 axoneme, 336-53 central sheath, 349 coarse fibres, 350-2 links heads, 349-50

Sperm cells-cont. axoneme-con t. matrix, 352-3 microtubdes, 338-49 cell surface, 3 17-24 centriolar region, 332-6 double flagellar apparatus, 367-74 mitochondria, 354-63 motility, 374-82 nucleus, 328-32 polymorphism and genetics, 382-3 Sphingomyelin, and choline metabolism, 53-5, 72-6, 83-4,92 Steatococcus tuberculatus, non-flagellate sperm, 370 Stegobium panicea, choline in development, 56 Stenodema calcaratum, salivary pectinase, 2 13 Sternorrhyncha, saliva, 192, 225 Stick insect, fat body deposits, 260 Stomacoccus plantani, non-flagellate sperm, 370 Strauzia longipennis, lipids containing choline, 73 Strontium and stimulation by ADH, 40 Structure-activity relationships, 5-HT, 5-12 Strychnine sulphate, and learning, 169 Stylet-sheath feeding, Hemiptera discharge of saliva, 197-200 formation of sheath, 196-7 ingestion, 200-1 and phytopathogenicity, 21 7 sampling the surface, 194-5 secretion of flange, 195-6 sheath material, composition, 205-8 withdrawal of stylets, 201-2 Subcellular fractions, lipids containing choline, 76 Suboesophageal ganglion, and learning, 136 Succinic dehydrogenase, sperm mitochondria, 364 Sucrose-gap technique, 22, 277, 284 Sulphydryl groups, Hemipteran saliva, 231,240,245,246 Symbionts, and choline metabolism, 58, 59

SUBJECT INDEX

Sympathetic stimulation, salivary glands, 37 Synapse, reactions at, 96-7 Synaptic changes, during learning, 162 Synaptic transmission, and cyclic AMP and Calcium post-synaptic, 35-6 pre-synaptic, 34-5

T TEA, and potentials, 278, 283, 284, 286 Telamona, sperm axoneme, 342 Temperature, and sperm motility, 381 Tenebrio molitor choline metabolism lipids containing choline, 72 nutritional requirements, 55, 56, 59, 60, 63 phosphatidylcholine, 78, 8 1 sperm accessory flagellar bodies, 364, 365 acrosomal complex, 324, 325 axoneme, 337,343,348 cell surface, 323 centriolar region, 336 mitochondria, 358,361,362 motility, 377, 378 Tenebrionid beetles, carnitine, 52 TEPP (tetraethylpyrophosphate) and acetylcholine, 99, 100 Termites, sperm, 354, 37 1 Termopsidae, sperm, 37 1 Tetrodotoxin, and action potential, 278 Tettigonioid Orthoptera, sperm, 326 Thelmatoscopus albipunctatus, nonflagellate sperm, 372 Theophylline, and cyclic AMP, 14, 15, 18,27, 31, 34,38, 39 Thomasiniellula populicola, salivary pectinase, 2 13 Thorax tissues, choline, 75 Threonine, aphid saliva, 218 Thrips, sperm, 370

433

Thyroid stimulating hormone, and cyclic AMP, 37 Thysan optera saliva, 189, 191, 192,217 sperm, 327,338,351,355,369 Thysanura lipids containing choline, 7 1,74 sperm accessory flagellar bodies, 363, 364 acrosomal complex, 324, 327 axoneme, 349,351 mitochondria, 354 nucleus, 328 paired sperm, 367-9 “Tight” junctions, perineurial, 95, 285, 287, 290, 291, 300, 302, 304 . Timarchia tenebriosa, ionic composition of haemolymph, 275,276 Tineola bisselliella, choline in development, 57 Tingidae, feeding, 191, 192, 193, 203, and phytopathogenicity, 21 7 Toxoptera aurantii, B vitamins in development, 58 Transplantation of ganglia, cockroach, 150 Transporting epithelia, hormone regulation, 39-41 Trehalose from glycogen, hormones, 32 Trial-and-error learning, 113-5, 157-62, 164 Triatoma protracta, saliva, 205 Tribolium castaneum, choline substitutes, 59 Tribolium confusum, choline metabolism enzymes, 8 9 , 9 0 lipid-soluble metabolites, 72, 78,83 nutritional requirements, 56,59, 60 Trichoplusia, sperm capacitation, 38 1 Trichoptera, sperm acrosomal complex, 327, 328 axoneme, 338, 339, 342, 343, 346, 347,348 mitochondria, 355 spermatids, 370

434

SUBJECT INDEX

Triglycerides, and cyclic AMP, 38 Tris ions, and potential changes, 283, 284,285 Trogoderma granarium, choline metabolism, 5 6 , 7 1 , 7 2 , 7 8 Tryptamine, salivary gland stimulation, 7 Tryptophane Hemipteran saliva, 221, 223,224 salivary gland stimulation, 7 Tubulin, sperm, 341,343,349 Type R learning, 113-5, 157-62, 164 Type I1 learning, 1 13-5, 157-62, 164 Typhlocybidae, salivary glands, 230 Tyrosine, aphid saliva, 218

Vasopressin (ADH), and cyclic AMP, 39-40 Vasotocin, and cyclic AMP, 39 Vespa crabro, venom, acetylcholine, 66,92 Vicia faba, saliva, 189 Virus transmission, and saliva, 242, 244,250 Visceral muscles, hormone regulation, 32 Viscosity, and sperm motility, 38 1 Viteus vitifolii, saliva composition, 2 16 glands, 228-9 metabolites, 2 18-9 origin, 245 and phytopathogenicity, 21 7 , 220, 22 1

U “Under-asparagine”, aphid saliva, 2 18 Uterus, and cyclic AMP, 35 UTPase, sperm axoneme, 346, 349, 351,352,365,367

V Valine, Hemipteran saliva, 218,221

W Water transport, toad bladder, 39-40 Watery saliva, composition and function, 208-17

X Xenophyes cascus, salivary glands, 231

Cumulative List of Authors Numbers in bold face indicate the volume number o f the series Aidley, D. J., 4, 1 Andersen, Sven Olav, 2, 1 Asahina, E., 6, 1 Ashburner, Michael, 7, 1 Baccetti, Baccio, 9, 3 15 Beament, J. W. L., 2, 67 Berridge, Michael J., 9, 1 Boistel, J., 5, 1 Bridges, R. G., 9, 5 1 Burkhardt, Dietrich, 2, 131 Bursell, E., 4, 33 Burtt, E. T., 3, 1 Carlson, A. D., 6, 5 1 Catton, W.T., 3, 1 Chen, P. S., 3, 53 Colhoun, E. H., 1, 1 Cottrell, C. B., 2, 175 Dadd, R. H., 1 , 4 7 Dagan, D., 8, 96 Davey, K. G., 2,219 Edwards, John S., 6,97 Eisenstein, E. M., 9, 11 1 Fraser Rowell, C. H., 8, 146 Gilbert, Lawrence, I., 4, 69 Goodman, Lesley, 7 , 9 7 Harmsen, Rudolf, 6,139 Harvey, W. R., 3, 133 Haskell, J. A., 3, 133

Hinton, H. E., 5,65 Hoyle, Graham, 7, 349 Kilby, B. A., 1, 11 1 Lawrence, Peter A., 7, 197 Lees, A. D., 3, 207 Maddrell, S. H. P., 8, 200 Miles, P. W., 9, 183 Miller, P. L;, 3, 279 Narahashi, Toshio, 1, 175; 8, 1 Neville, A. C., 4, 213 Parnas, I., 8, 96 Pichon, Y.,9, 257 Prince, William T., 9, 1 Pringle, J. W. S., 5, 163 Rudall, K. M., 1,257 Sacktor, Bertram, 7,268 Shaw, J., 1,315 Smith, D. S., 1,401 Stobbart, R. H., 1, 315 Treherne, J. E., 1,401; 9, 257 Usherwood, P. N. R., 6,205 Waldbauer, G. P., 5,229 Weis-Fogh, Torkel, 2, 1 Wigglesworth, V. B., 2, 247 Wilson, Donald M., 5,289 Wyatt, G. R., 4, 287 Ziegler, Irmgard, 6,139

435


Cumulative List of Chapter Titles Numbers in bold face indicate the volume number of the series Active Transport and Passive Movement of Water in Insects, 2,67 Amino Acid and Protein Metabolism in Insect Development, 3, 53 Biochemistry of Sugars and Polysaccharides in Insects, 4, 287 Biochemistry of the Insect Fat Body, 1, 11 1 Biology of Pteridines in Insects, 6,139 Cellular Mechanisms Underlying Behaviour-Neuroethology, 7,349 Chitin Orientation in Cuticle and its Control, 4, 213 Chitin/Protein Complexes of Insect Cuticles, 1, 257 Choline Metabolism in Insects, 9, 5 1 Colour Discrimination in Insects, 2, 131 Comparative Physiology of the Flight Motor, 5, 163 Consumption and Utilization of Food by Insects, 5, 229 Control of Polymorphism in Aphids, 3, 207 Control of Visceral Muscles in Insects, 2, 2 19 Effects of Insecticides on Excitable Tissues, 8, 1 Electrochemistry of Insect Muscle, 6,205 Excitation of Insect Skeletal Muscles, 4, 1 Excretion of Nitrogen in Insects, 4, 33 Feeding Behaviour and Nutrition in Grasshoppers and Locusts, 1 , 4 7 Frost resistance in Insects, 6,1 Function and Structure of Polytene Chromosomes During Insect Development, 7, 1 Functional Aspects of the Organization of the Insect Nervous System, 1,401 Functional Organizations of Giant Axons in the Central Nervous Systems of Insects: New Aspects, 8, 96 Hormonal Regulation of Growth and Reproduction in Insects, 2, 247 Image Formation and Sensory Transmission in the Compound Eye, 3, 1 Insect Blood-Brain Barrier, 9, 257 Insect Ecdysis with Particular Emphasis on Cuticular Hardening and Darkening, 2, 175 Insect Sperm Cells, 9 , 315 Learning and Memory in Isolated Insect Ganglia, 9 , 11 1 Lipid Metabolism and Function in Insects, 4, 69 Mechanisms of Insect Excretory Systems, 8, 200 Metabolic Control Mechanisms in Insects, 3, 133 Nervous Control of Insect Flight and Related Behaviour, 5,289 Neural Control of Firefly Luminescence, 6,5 1 Osmotic and Ionic Regulation in Insects, 1, 3 15 '

431

438

CUMULATIVE LIST OF CHAPTER TITLES

Physiological Significance of Acetylcholine in Insects and Observations upon other Pharmacologically Active Substances, 1, 1 Polarity and Patterns in the Postembryonic Development of Insects, 7 , 197 Postembryonic Development and Regeneration of the Insect Nervous System, 6, 97 Properties of Insect Axons, 1, 175 Regulation of Breathing in Insects, 3, 279 Regulation of Intermediary Metabolism, with Special Reference to the Control Mechanisms in Insect Muscle, 7 , 2 6 8 Resilin. A Rubberlike Protein in Arthropod Cuticle, 2, 1 Role of Cyclic AMP and Calcium in Hormone Action, 9, 1 Saliva of Hemiptera, 9 , 183 Spiracular Gills, 5, 65 Structure and Function of the Insect Dorsal Ocellus, 7 , 9 7 Synaptic Transmission and Related Phenomena in Insects, 5, 1 Variable Coloration of the Acridoid Grasshoppers, 8, 146

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