ADVANCES IN INSECT PHYSIOLOGY VOL 4 APL, Volume 4 (v. 4)

Advances in

INSECT PHYSIOLOGY

VOLUME 4

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Advances in

Insect Physiology Edited by

J. W. L. BEAMENT, J. E. TREHERNE and V. B. WIGGLESWORTH Department of Zoology, The University, Cambridge, England

VOLUME 4

1967

ACADEMIC PRESS London and New York

ACADEMIC PRESS INC. (LONDON) LTD BERKELEY SQUARE HOUSE BERKELEY SQUARE LONDON, W.1

US.Edition published by ACADEMIC PRESS INC.

11 1 FIFTH AVENUE 10003

NEW YORK, NEW YORK

Copyright 0 1967 By Academic Press Inc. (London) Ltd

All Rights Reserved NO PART OF THIS BOOK MAY BE REPRODUCED IN ANY FORM, BY PHOTOSTAT, MICROFILM, OR ANY OTHER MEANS, WITHOUT WRITTEN PERMISSION FROM THE PUBLISHERS

Library of Congress Catalog Card Number: 63-14039

Printed in Great Britain by William Clowes and Sons, Limited, London and Beccles

Contributors to Volume 4 D. J. AIDLEY, Department of Zoology, University of Oxford, England (p. 1)

E. BURSELL,Department of Biological Sciences, University of Rhodesia and Nyasaland, Salisbury, Rhodesia @. 33) L. I. GILBERT, Department of Biological Sciences, Northwestern University, Evanston, Illinois, U.S.A. (p. 69) A. C . NEVILLE, Department of Zoology, University of Oxford, England (p. 213) G. R. WYATT, Department of Biology, Yale University, New Haven, Connecticut, U.S.A. (p. 287)

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Contents CONTRIBUTORSTO VOLUME4

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V

EXCITATION OF INSECT

SKELETAL. D. J. AIDLEY

I. Introduction

.

11. The Resting Potential

MUSCLES

.

A. The effect of potassium ions . B. The effect of chloride ions . C. The effect of sodium ions . D. The effect of divalent cations . E. The effects of carbon dioxide F. Temperature dependence . 111. Neuromuscular Transmission . A. The innervation of insect muscle . B. Excitatory responses . C. The quanta1 release of transmitter substance . D. Inhibitory responses . , IV. The Electrical Excitability of the Muscle Fibre Membrane A. The electrical properties of electrically excitable responses B. The ionic basis of electrically excitable responses . C. Spontaneous activity . . V. The Excitation-contraction Coupling Process . A. Depolarization . B. The importance of calcium ions . C. The action of carbon dioxide . D, “Fast” and “slow” muscles . References .

.

1 2 2 4 5 6 6 6 7 7 8 15 17 20 20 21 23 23 23 24 26 26 27

THEEXCRETION OF NITROGEN IN INSECTS E. BURSELL

I. Introduction . 11. The Formation of Nitrogenous End Products A. The uricolytic pathway . . B. The uricotelic pathway . C. The formation of urea . D. The formation of ammonia . E. Amino acids . F. Miscellaneous materials . III. The Excretion of Nitrogenous End Products A. Collembola . B. Orthoptera . C. Odonata .

vii

.

33 34 36

40

.

41 42 43 44 44 45 46

47

viii

CONTENTS

D. Dermaptera

E. Hemiptera . F. Coleoptera G. Neuroptera . H. Hymenoptera I. Diptera . J. Lepidoptera 1V. Conclusions . References .

.

48 48 50 51 51 52 54 56 61

.

. .

LIPIDMETABOLISM AND FUNCTION IN INSECTS LAWRENCE I. GILBERT

I. Introduction . A. General . B. Definition and classification . 11. Lipid Content . A. Expression of data . B. Alterations during metamorphosis . C. Nature of insect lipids . III. Lipid Utilization . A. Digestion and absorption . B. Lipid release and transport . C. Extra-digestive lipases . D. Fatty acid catabolism . IV. Lipid Biosynthesis . A. General mechanism of fatty acid synthesis B. Fatty acid biosynthesis in insects . C. Phospholipid and triglyceride . D. Fatty acids in nutrition . E. Substrate interconversion . V. Hydrocarbons and Waxes . A. Cuticle B. Extra-cuticular . VI. Isoprenoid Compounds . A. Nutritional studies . B. Isoprenoid biosynthesis . C. Isoprenoid content . D. Sterol modification . E. Function F. Insect hormones . W.Conclusions . References . Addenda .

70 70 71 71 71 81 89 97 97 102 110 116 127 127 130 134 145 147 152 152 155 157 157 161 168 170 175 176 186 187 208

.

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.

CHITIN ORIENTATION IN CUTICLE AND

ITS

CONTROL

A. C. NEVILLE

I. Introduction

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II. Orientation and the Mechanicai Properties. of &tic& molecules

.

213

Macro:

217

CONTENTS

ix

. A. Parallel orientation . B. Crossed fibrillar orientation . C. Lamellar structure . D. Functional aspects . 1V. Orientation Control . A. Circadian organization . B. Metabolic oscillators and “switches” C. “Dermal” light sense . D. Implantation experiments . E. Nervous control . F. Discussion . V. Orientation Mechanisms . A. Primary orientation . B. Secondary orientation . C. Protein orientation . D. Hypothesis . M.Conclusion . References .

220 220 220 223 229 233 233 246 254 257 260 260 262 263 265 270 272 279 280

III. Types of Chitin Architecture

THEBIOCHEMISTRY OF SUGARS AND POLYSACCHARIDES IN INSECTS G. R. WYATT

1. Introduction . II. The Occurrence of Sugars in Insects . A. Glucose and reducing substances . B. Trehalose . C. Sugar content of insect hsmolymph . D. Sugar content of whole insects and insect tissues . 111. Intestinal Absorption and the Physiology of Hemolymph Sugar Levels . A. Absorption from the gut . B. Regulation of blood sugar . IV. Biosynthesis and Utilization of Sugars . A. Glucose . B. Use of monosaccharides other than glucose . C. Biosynthesis of trehalose . D. Cleavage and use of trehalose . E. Physiological roles of trehalose and trehalase . F. Dormancy and the properties of trehalose . .* . V. Glycogen . A. Glycogen in insects . B. Accumulation and conversion during growth and metamorphosis C. Glycogen in insect flight D. Metabolism of glycogen VI. Hormonal Effects on Carbohydrate Metabolism .

.

1.

. .

287 289 289 289 29 1 295 297 297 299 301 301 302 304 309 317 324 325 325 327 329 329 336

CONTENTS

X

W.Glycoproteins and Chitin

.

A. Glycoproteins in insects B. Metabolism of chitin . Vm. Glycerol and Sorbitol . . References

AUTHORINDEX .

SUBJECT INDEX

.

.

.

. .

340 340 341 345 347

.

361

. .

. 375

The Excitation of Insect Skeletal Muscles D. J. AIDLEY Department of Zoology, University of Oxford, England I. Introduction

.

11. The Resting Potential

1

.

A.

The effect of potassium ions B. The effect of chloride ions . C. The effect of sodium ions . D. The effect of divalent cations . E. The effects of carbon dioxide . F. Temperature dependence . UI. Neuromuscular Transmission . A. The innervation of insect muscle . B. Excitatory respomes . C. The quanta1 release of transmitter substance . D. Inhibitory responses . IV. The Electrical Excitability of the Muscle Fibre Membrane . A. The electrical properties of electrically excitable responses B. The ionic basis of electrically excited responses . C. Spontaneous activity . . V. The Excitation-contraction Coupling Process A. Depolarization . B. The importance of calcium ions . C. The action of carbon dioxide . D. “Fast” and “slow ” muscles References .

. . .

. . . . . . .

. . . . . . .

.

2 2 4

s

6 6 6 7 7 8 1s 17 20 20 21 23 23 23

. 2 4 26 . 26 . 27

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I. INTRODUCTION

In the resting condition, muscle cell membranes ‘are electrically polarized so that the inside of the cell is a few tens of millivolts negative to the outside. Stimulation of the motor nerves supplying the muscle results in a reduction of this membrane potential, which is followed by contraction of the muscle. This article will attempt to view our knowledge of this chain of events in insect muscles mainly in the light of the much greater understanding of vertebrate muscles. I

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D . J . AIDLEY

11. THERESTINGPOTENTIAL A . THE EFFECT OF POTASSIUM IONS

According to the theory developed by Boyle and Conway (1941) to explain the ionic distribution between the cell and the extracellular fluid in frog sartorius muscle, the distribution of potassium and chloride ions follows a Donnan equilibrium set up by the presence of indiffusible anions inside the fibre, the system being maintained in osmotic equilibrium by the presence of an effectively indiffusible cation (sodium) in the extracellular fluid. The resting potential at equilibrium (E) is then given by the equations

where R is the gas constant, T the absolute temperature, F is Faraday’s constant, and the subscripts and ,refer to the intracellular and extracellular media, respectively. These equations also define the potassium equilibrium potential, EK,and the chloride equilibrium potential, Eel. Changes in [Kl0 or [CI], result in movements of KCl across the membranes so as to restore the equilibrium condition given by eqs. (1) and (2). Since movement of a given quantity of KCI results in much greater relative changes in [CI], than in [K],, changes in [Kl0 cause much larger changes in membrane potential at equilibrium than do changes in [Cl],, although large transient changes in membrane potential may be produced by changes in [Cl], (Hodgkin and Horowicz, 1959). One method of testing the applicability of eq. (1) is to measure [K], and [K], and see if the membrane potential calculated from the equation agrees with that actually found. This was done by Wood (1963) for leg muscle fibres of Locusta, Periplaneta and Carausius after equilibration for four to six hours in salines of ionic composition approximating to that of the haemolymph, with [K], between 10 and 18 m ~ In. each case the actual resting potential was close to that predicted from eq. (1). A more usual method of testing the applicability of eq. (1) is by measuring membrane potentials at different values of [K],. It is necessary either to measure [K], at each value of [Kl0 (Conway, 1957), or to ensure that there is no change in [K], during the course of the experiment by maintaining the [K],[Cl], product constant or by eliminating chloride from the system by using extracellular solutions

THE EXCITATION O F INSECT SKELETAL MUSCLES

3

containing sulphate as the major anion (Adrian, 1956; Hodgkin and Horowicz, 1959). If [K]*is constant, eq. (1) predicts that the membrane potential should be related to [K], by means of a straight line with a slope (at 18") of 58 mV per unit change in log,,[K],. Hoyle (1953) studied the effects of substitution of potassium chloride for sodium chloride in the external saline solution on the membrane potential of fibres in the metathoracic retractor unguis of Schistocercu (Fig. 1). At concentrations greater than 10 mM, the relation between

Potassium concentration (mu)

RQ.1. The relation between potassium ion concentration and membrane potential in the metathoracic retracto: unguis muscle of Schisrocerca. Chloride ion concentration constant. (From Hoyle, 1953.)

membrane potential and log,,[K], was linear, with a slope of about 50 mV. Similar results have been obtained by Hagiwara and Watanabe (1954) from locust flight muscle (slope 25 mV) and Wood (1957) on the prothoracic flexor tibialis of Curuusius (slope 36 mv). In each case the slope of the straight line section of the curve was less than the 58 mV predicted by eq. (1). However, the method used in these experiments involves a change in the [K],[CI], product so" that the system will initially be displaced from equilibrium (so that EE is not equal to Ec3

4

D . J . AIDLEY

and can only reach a new equilibrium position by a change in [K],; both these effects will tend to reduce the slope of the line. When the external potassium ion concentration is low, the membrane potential is less than that predicted from eq. (I), so that the graph of membrane potential against log,,[K], tends to level out in this region (Fig. 1). This phenomenon is common to a number of types of cell, and can be ascribed to a constant small permeability of the membrane to sodium ions and a decrease in permeability to potassiuni ions when [K], is low (Hodgkin, 1951). The membrane potential can then be described by the equation

(Hodgkin and Katz, 1949), where PK,PNaand PClare permeability coefficients. There seems to be little doubt that this equation could be successfully applied to the results obtained from the experiments so far mentioned. There are, however, observations on some insects which are very difficult to reconcile with the potassium electrode hypothesis for the resting potential. Belton (1958, 1960) reported that the membrane potentials of muscle fibres of various Lepidoptera were unaffected by changes in [K],. Belton and Grundfest (1962) obtained similar results from Tenebrio larvae; here the membrane potential was not affected by changes in [K],up to about 100 m, but at higher concentrations there was a logarithmic relation between membrane potential and [Kl0with a slope of 58 mV. They also measured the potassium concentration of the muscle fibres (72 m) and the haemolymph (40 m),and pointed out that the value of EKcalculated from these figures (- 17 mv) is much less than the actual membrane potential (- 50 to - 60 mV in muscles bathed in haemolymph). B . T H E EFFECT O F C H L O R I D E IONS

The relationship between external chloride ion concentration and the membrane potential has been investigated by Wood (1965), in the leg muscle fibres ofbcusta and Periplaneta. In both insects there was an increase of a few millivoltsas [CI], was raised from 0 to 150mM. This implies that the membrane is permeable to some extent to chloride ions. At first sight, the fact that the membrane potential was not related to [Cl], by a straight line with a slope of 58 mV per unit change in logl0[C1], might be explained by movement of KCl out of the cell when [Cl], was less than about 100 mM. However, Wood found that [Cl], was almost independent

THE EXCITATION OF INSECT SKELETAL MUSCLES

5

of [Cl], in this range. He suggested that the results implied that the muscle

cell membrane is much less permeable to chloride than to potassium ions, with the possibility that there is some active transport of chloride across the membrane. Thus the insect muscle cell membrane would seem to be, in these respects, similar to the squid giant axon (Keynes, 1963) rather than the frog muscle fibre membrane, where the chloride permeability is high (Hodgkin and Horowicz, 1959). The importance of such a relatively low chloride permeability might be that it allows the membrane potential to be changed by increase of PCl during inhibition (see Section 111, D, below). However, Wood’s results cannot be taken as implying that the chloride conductance of the membrane is negligible. If this were so, it would be difficult to explain the departure of the slope of the relation between membrane potential and log,,[K], from the 58 mV predicted by eq. (1). A critical investigation of this problem would include measurements of the transient changes in membrane potential caused by changes in [Kl0 and [Cl],, such as were made by Hodgkin and Horowicz (1959) on frog muscle fibres. Usherwood and Grundfest (1965) noticed that transient depolarizations of 7 to 15 mV occurred when a chloride-free solution was substituted for the normal Ringer solution perfusing Romalea muscle fibres ;the reverse procedure caused transient hyperpolarizations (Usherwood, personal communication). C . THE EFFECT OF S O D I U M IONS

Wood (1961, 1963) showed that a decrease in external sodium ion concentrationresulted in a fall in the resting potential of muscle fibres of Lmusta, Periplaneta and Carausius. The tonicity of the low-sodium salines was maintained by substitution of sucrose or choline chloride for sodium chloride. In crustacean muscle fibres, Fatt and Katz (1953) observed a similar decrease in the resting potential when sucrose was substituted for sodium, but only a very small decrease when sodium was replaced by choline. In the case of sucrose substitution it is evident that the associated decrease in chloride concentration (and, probably, consequent movements of KC1) would be expected to lower the membrane potential, although it is not clear whether these effects would be sufficient to account for the magnitude of the observed changes. Obviously this explanation cannot be applied to the fall in membrane potential which occurs in insect muscles when choline chloride is used as a substitute for sodium chloride. Hence it seems (Wood, 1961) that there is some direct influence of sodium ions on the permeability of the resting muscle fibre membrane.

6

D. J. AIDLEY

There is at the time of writing no evidence for the existence of an active sodium extrusion mechanism in insect muscle fibres, but it would be most surprising if such a mechanism did not exist. D . T H E EFFECT O F D I V A L E N T C A T I O N S

It is well known that, in vertebrate muscles, decrease in the calcium ion concentration of the Ringer's solution causes a fall in resting potential, and that this fall can be prevented by the presence of other divalent cations (Jenden and Reger, 1963). This phenomenon has apparently not been looked for in insect muscles, but Wood (1957) showed that an increase in external calcium ion concentration caused an increase in the resting potential of Carausius muscle fibres ; increase in magnesium ion concentration did not affect the resting potential. High concentrations of barium ions cause a fall in the resting potentials of Romalea muscle fibres (Werman, McCann and Grundfest, 1961). E . T H E EFFECTS O F C A R B O N D I O X I D E

In Schistocerca spiracular muscle, introduction of carbon dioxide into the tracheal system causes a slight depolarization of the muscle fibre membranes, associated with a decrease in the membrane resistance (Hoyle, 1960). The ionic basis of this depolarization is not known, but an increase in sodium permeability seems to be the obvious candidate. Similar slight depolarization by carbon dioxide occurs in the flight muscles of the beetles Pissodes and Tenebrio, but very large depolarizations are produced in the flight muscles of the wasp Vespula and the fly Sarcophaga (McCann and Boettiger, 1961). In the pupal spiracular muscles of the silkmoths Hyalophora and Telea, application of carbon dioxide causes a hyperpolarization of the fibre membrane (van der Kloot, 1963). F . TEMPERATURE DEPENDENCE

Kerkut and Ridge (1 96 1) compared the immediate effects of a change in temperature from 15 to 25" on the resting potentials of muscle fibres of Carcinus, Rana and Periplaneta. The Qlo values over this range were 1.064 & 0.004(s.E.)for the crab, 1.060 2 0.004 for the frog, and 1.296 f 0.046 for the cockroach. These values are all significantly different from the value of 1.035 expected if the system obeys eq. (I), and the value for the cockroach is significantly higher than those for the frog and the crab. Kerkut and Ridge concluded that the resting potential is directly dependent upon metabolic processes. Since the measurements of membrane

THE EXCITATION O F I N S E C T S K E L E T A L MUSCLES

7

potential were made after about 1 min. at the new temperature, it is improbable that this effect is dependent upon changes in ionic concentrations in the cell. If there is a sodium extrusion mechanism which is not electrically neutral (for example, if there is no 1 :1 exchange with potassium ions) the resting potential will be dependent to some extent on the activity of the pump, being more negative when the pump is active (as it presumably is at higher temperatures within the physiological range). However, this is not the only possible interpretation of these results. As we have seen, eq. (3) gives a better description of the resting potential than does eq. (l), particularly at external potassium ion concentrations less than about 10 mM; in the salines used by Kerkut and Ridge the potassium ion concentration was approximately 5 mM. Now the Qlo of membrane potential given by eq. (3) will only be 1.035 for the range 15 to 25” if the Qlos of the permeability coefficients PK,P,, and Pcl are identical. In particular, if the Qlo of PNais less than those of PK and Pel, increase in temperature will result in a hyperpolarization greater than that predicted by eq. (1). It should be possible to distinguish between these two alternatives by measuring the effect of temperature changes on the membrane potential of fibres poisoned with some suitable metabolic inhibitor. 111. NEUROMUSCULAR TRANSMISSION A. THE I N N E R V A T I O N O F I N S E C T MUSCLE

This subject has been extensively discussed in a recent review by Hoyle (1965), and will therefore be but briefly examined here. Insect muscle fibres possess multiterminal innervation, there being numerous motor nerve endings on each fibre which are spaced at intervals of 30 to 80 p apart along the length of the fibre. Polyneuronal innervation also occurs, so that many muscle fibres are innervated by more than one motor axon. In these cases stimulation of the different motor axons causes different responses in the muscle fibre. The motor axons can be divided into two broad categories: excitor axons, which produce depolarization of the muscle fibre membrane leading to contraction of the fibre, and inhibitor axons, which produce hyperpolarization and can counteract, to some extent, the action of the excitor axons. In many cases fibres are innervated by more than one excitor axon, and stimulation of these axons produces electrical and mechanical responses of different magnitude; axons producing larger responses are then known as “fast” axons, and those producing,smallerresponses are known as ‘“slow” axons (Pringle, 1939; Hoyle, 1955a, b). Some muscles (such as the extensor tibiae of

8

D . J . AIDLEY

locusts) are innervated by only one set of axons, while others (such as the flexor tibiae of locusts) are innervated by a number of axons of each type. The latter may thus be said to possess a number of motor units, although it is obvious that the definition of a motor unit in a system in which there may be overlapping fields of motor innervation, between axons producing different types of response, is largely a matter of opinion. These points may be illustrated by reference to two muscles of the locust Schistocerca gregaria, in which the pattern of motor innervation has been closely examined. The metathoracic extensor tibiae (the jump ing muscle) is innervated by three motor axons (Hoyle, 1955a, b), a “fast” excitor, a “slow” excitor and an inhibitor. About 80% of the muscle fibres are innervated by the “fast” exciter, 20 to 30% are innervated by the “slow” excitor, and about 10% (all of them also innervated by the ‘‘slow’’ excitor) are innervated by the inhibitor (Usherwood and Grundfest, 1965). Thus some fibres are innervated by all three axons, some are innervated by the “ slow” excitor and either the inhibitor or the “fast” excitor, and the majority are innervated by the “fast” excitor only. In the meso- and metathoracic dorsal longitudinal flight muscles of Schistocerca, there are five motor units each separately supplied by one axon, the response to stimulation being of the “fast” type (Neville, 1963). It is probable that there is no “slow” or inhibitor innervation of these muscles. B. EXCITATORY RESPONSES

1. The general nature of the excitatory responses Stimulation of “slow” type excitatory axons causes a depolarization of the muscle fibre membrane, the postsynaptic potential (also known as the junction potential or end plate potential), which rises fairly rapidly to a peak and then falls more slowly (Fig. 2). The size of the response varies between different preparations and between different fibres of the same preparation; Hoyle (1957) gives a range of 2 to 50 mV for the size of the “slow” postsynaptic potential in the mesothoracic extensor tibiae of Schistocerca. Stimulation of “fast” type excitatory axons causes a larger electrical response (Fig. 3), which frequently overshoots the zero level of membrane potential so that the inside of the fibre becomes briefly positive to the outside. In many cases the “fast” response consists of two components, a postsynaptic potential and an electrically excited response

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9

(sometimes called the “active membrane response”) caused by the depolarization constituting the postsynaptic potential (see Section IV below). This electrically excited component may be absent in some preparations, such as the metathoracic spiracular muscle of Schistocerca (Hoyle, 1959). There does not seem to be any fundamental difference between “slow” and “fast” responses. “ Slow” responses may summateto give a depolarization large enough to elicit an electrically excited response (Cerf et al., 1959),and “fast” responses lose their regenerativecomponents and look

FIG.2. The electrical response to stimulation of the “fast” axon in the mesothoracic extensor tibiae of Schistocercu. Intracellular records from three different fibres. The upper trace shows zero potential initially, then twitch tension.

just like “slow” responses when the muscle is subjected to low temperatures (del Castillo et al., 1953) or neuromuscular blocking agents (Hill and Usherwood, 1961; Hoyle, 1955~). 2. The ionic basis of excitatory postsynaptic potentials In their classic study of the postsynaptic potential in frog sartorius muscle, Fatt and Katz (1951) suggested that the chemical transmitter substance (acetylcholine in this case) released from the motor nerve ending causes a brief increase in the ionic permeability of the postsynaptic membrane, so that the resting potential is effectively shortcircuited at the motor end plate during the action of the transmitter

10

D. J . A I D L E Y

substance. In accordance with this hypothesis, it was found that the size of the postsynaptic potential increased proportionately when the membrane potential was increased by passing current through a second microelectrode inserted in the (curarized) muscle fibre. A similar effect

. . c-w-!=Qc

I 100 rnsec

FIG.3. Theelectrical response to stimulation of the “slow” axon in the mesothoracic extensor tibiae of Schistocerca. Intracellular records from four different fibres (a-d). Upper trace shows zero potential and stimulus monitor. Note reflex responses in c and summation in the right-hand record from ‘0.

was found by del Castillo et al. (1953) for the “fast” responses of locust leg muscles, the postsynaptic potential component of the electrical response being just over half the value of the membrane potential over the range - 50 to - 150 mV. Further confirmation of this view was provided by Cerf et al. (1959), who showed that depolarizations large enough to make the inside of the fibre positive to the outside resulted in a

THE EXCITATION OF INSECT SKELETAL MUSCLES

11

reversal of the sign of the “slow” postsynaptic potential. Similarly, depolarization of the muscle fibre membrane by increasing the external potassium ion concentration causes a decrease in the size of the postsynaptic potential (Hoyle, 1955~;Wood, 1957). In frog muscles, voltage clamp studies have shown that sodium and potassium are the ions mainly involved in the increase in permeability during the postsynaptic potential (Takeuchi and Takeuchi, 1960). There is as yet no information of this kind for the excitatory postsynaptic potentials of insect muscles.

3. The blocking effect of tryptamine and similar compounds Hill and Usherwood (1961) showed that perfusion of the metathoracic flexor and extensor tibiae muscles of Schistocerca with solutions containing 1 to 10 DIM tryptamine diminished both the electrical and

FIG.4. The effect of tryptamine on the “fast” response of the metathoracicflexor tibiae of Schistocerca. The upper trace shows tension and, initially, zero potential. Time signal 500 c/s. Trace 1 shows the normal response. Traces 2-4 show responses at 5 sec. intervals after the application of 10 mM tryptamine. Traces 6-10 show responses at 5 sec. intervals after removal of tryptamine. (From Hill and Usherwood, 1961.)

mechanical responses to “fast” axon stimulation (Fig. 4). Similar results were obtained with 5-hydroxytryptamine, lysergic acid diethylamide, bromolysergic acid diethylamide, 5,6-dimethoxytryptamine and 3-(pyrrolidino-methyl)-thionapthene. These substances did not affect the conduction of action potentials in the crural nerve trunk, and neither tryptamine nor 5-hydroxytryptamineaffected the muscle fibre membrane resistance or the electrically excited responses of the membrane to depolarization via an internal electrode. It was concluded that these substances act as neuromuscular blocking agents, possibly by competing with the transmitter substance for receptor sites 04 the postsynaptic membrane.

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D. J . A I D L E Y

Hill and Usherwood also found that tryptamine increases the synaptic delay, and that this effect appears to be more long-lasting than the blocking effect on the size of the post-synaptic potential. In the mesothoracic extensor tibiae of Schistocerca tryptamine also blocks transmission from the “slow” axon (D. J. Aidley, unpublished). 4. The nature of the excitatory transmitter substance A number of studies have been made on the effects of acetylcholine on

insect muscles. Harlow (1958) found that acetylcholine had no effect on resting locust leg muscles, and Wood (1958) obtained similar results for the prothoracic flexor tibiae of Carausius. These authors also reported that d-tubocurarine and adrenaline did not affect the responses to nervous stimulation. On the other hand, it seems that neurally evoked contractions can be increased in size in the presence of acetylcholine. This effect has been described by Harlow (1958) and Hill (1963) in locusts, and by Kerkut et al., (1965a) in the cockroach. The nature of the mechanisms behind this increase in contraction size is obscure; there is an urgent need for microelectrode studies on the form of the electrical responses to stimulation under these conditions. Similar increase in the contraction size is brought about by L-glutamic acid and, to a lesser extent, D-glutamic and L-aspartic acids (Kerkut et al., 1965a). The presence of glutamate in perfusates of the femoral muscles of Periplaneta has recently been demonstrated by Kerkut et al. (1965b). They found that the amount of glutamate produced was proportional to the number of stimuli to the motor nerves, and that the muscles contracted when perfused with low concentrations of glutamate, the contraction increasing in size as the glutamate concentration was increased. Similar results were obtained for crab and snail muscles. Usherwood and Grundfest (1965) state that glutamate causes a depolarization of insect muscle fibres. It is evident from these results that a number of substances have excitatory effects on insect muscles, although it is impossible at present to say which of them, if any, is the naturally occurring excitatory transmitter substance. Glutamate, or a glutamate complex, appears to have the strongest claims in this respect. It is possible that the “slow” and “fast” responses to nervous stimulation are brought about by different transmitter substances, but it seems more likely that “fast” responses are brought about by release of more transmitter substance per impulse than are “slow” responses. Qualitatively, the effects of blocking agents appear to be the same on

T H E E X C I T A T I O N OF I N S E C T S K E L E T A L MUSCLES

13

both systems. This opinion would have to be revised if it were shown that the reversal potentials for the two types of postsynaptic potential were different. 5. Calcium-magnesium antagonism Increase of the magnesium ion concentration of the external solution causes a diminution in the size of the “fast” postsynaptic potential in Locusta and Periplaneta (Hoyle, 1955~).This effect is antagonized by a corresponding increase in calcium ion concentration. “ Slow ” postsynaptic potentials are similarly affected @. J. Aidley, unpublished). These results are similar to those obtained by del Castillo and Engbaek (1954) from frog sartorius muscles. They concluded that the depressant action of magnesium occurs presynaptically, part of the evidence for this being that the very small postsynaptic potentials produced in high magnesium ion concentrationsshow quanta1fluctuations in size, thought to correspond with “packets” of acetylcholine released from the motor nerve ending (del Castillo and Katz, 1954). Similar fluctuations in postsynaptic potential size were observed by Usherwood (1963a) in Schistocerca leg muscles in solutions containing high magnesium ion concentrations, and hence it would seem probable that the site of this calcium-magnesium antagonism is presynaptic in locust muscle. The blood of some herbivorous insects may contain relatively large concentrations of magnesium ions (Duchilteau et al., 1953); in Carausius, for example, investigated by Wood (1957), the average magnesium concentration was 53 mM, whereas that for calcium was only 7.5 m. This situation is of some interest, for concentrations of this order are sufficient to block completely neuromuscular transmission in locusts and frogs. Wood found that the size of the “fast” postsynaptic potential decreased as the calcium ion concentration was reduced below its value in the blood, as in locusts and frogs. However, the relation which he found between the size of the total electrical response to stimulation of the “fast” axon (postsynaptic potential plus electricallyexcited response) and the magnesium ion concentration is unique among those preparations which have been investigated. The electrical response increased from a value of about 21 mV in the absence of magnesium ions to a maximum of about 43 mV at a concentration of approximately 75 mM, thereafter falling with increasing magnesium ion concentration to reach zero at about 200 m. This fall in the electrical response at very high magnesium concentrations could not be antagonized by calcium ions, and may imply, as Wood suggested, that a high”magnesiumion concentration reduces the sensitivity of the postsynaptic membrane to the

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D . J. AIDLEY

transmitter substance, as appears to be the case in the frog sartorius muscle (del Castillo and Engbaek, 1954). It is not clear whether the increase in the size of the postsynaptic potential with increasing magnesium ion concentration over the range 0 to 75 mM is a presynaptic or a postsynaptic effect. 6. The action of carbon dioxide Postsynaptic potentials of Schistocerca metathoracic spiracular muscle (Hoyle, 1960) and the flight muscles of certain beetles and the bug Nazzara (McCann and Boettiger, 1961)are reduced in the presence of high concentrations of carbon dioxide. It is not clear whether the carbon dioxide acts presynaptically or postsynaptically in these cases.

7 . Time-dependentproperties of the postsynaptic potentials When the interval between motor nerve impulses is short enough, postsynaptic potentials readily summate so as to give a depolarization greater than that following a single impulse. In some cases, repetitive stimulation of the “slow” axon results in a “plateau” of depolarization whose degree is proportional to the stimulation frequency (Hoyle, 1955b; Wood, 1958). When the postsynaptic potential is accompanied by an electrically excited response (as in typical “fast” potentials) the situation is complicated by the presence of refractoriness in the electrically excited component, but it is sometimes possible to discern summation of the postsynaptic potential components when the stimulus interval is short enough (Wood, 1958). Some degree of facilitation is frequently present when the postsynaptic potentials are very small, but this phenomenon does not seem to be as well developed in insects as it is in crustacea. By estimating the quanta1 content of postsynaptic potentials in crustacean muscle fibres, Dude1 and Kuffler (1961) were able to show that facilitation is a presynaptic phenomenon, being caused by an increase in the amount of transmitter substance released from the motor nerve ending. It is probable that a similar mechanism occurs in insect muscles. Becht et al. (1960)claimed that, in some of the coxal muscles of the cockroach, the postsynaptic potential component of the second of two responses to “fast” axon stimulation showed the effects of refractoriness in that it was smaller than usual for about 15 msec. after the first response. The interpretation of this result is complicated by the presence of the electrically excited component associated with the first response, but it may well be that a process of depression (the reverse of facilitation) occurred.

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C. THE Q U A N T A L RELEASE OF TRANSMITTER SUBSTANCE

1. Spontaneous miniature postsynaptic potentials If a microelectrode is inserted into a frog sartorius muscle fibre in the region of the end plate, small “miniature postsynaptic potentials” can be seen (Fatt and Katz, 1952). These are similar in their pharmacology to neurally evoked postsynaptic potentials, and are probably caused by the spontaneous release of acetylcholine in “quantal” units derived from the vesicles which occur in the presynaptic nerve ending (Katz, 1962). Usherwood (1961, 1963a) has observed similar spontaneousminiature potentials in the muscle fibres of Schistocerca, Blaberus and Periplanetu (Fig. 5). The amplitude and time course of these miniature potentials were more variable than in the frog sartorius, an effect which is almost certainly due to the multiterminal innervation of insect muscles, so that potentials originate at different distances from the recording site, with consequent differences in the degree of attenuation of the potentials. In accordance with this view, Usherwood showed that the frequency distribution of the amplitudes of the potentials was much more skew in fibres which were long in relation to their length constant than in short ones. Analysis of the frequency distribution of the time intervals between successive potentials in a long series showed that the discharge is a random process. The average frequency of occurrence of the spontaneous miniature potentials could be increased by raising the external calcium ion concentration, and decreased by raising the external magnesium ion concentration. Concentrations of magnesium greater than 10 m~ also reduced the amplitude of the potentials; it is thought that this effect is by means of a reduction in the sensitivity of the postsynaptic membrane to the transmitter substance. Increase in external potassium ion concentration caused an increase in the average frequency of the potentials, probably by depolarizing the motor nerve endings. Application of salines made hypertonic with sucrose caused a transient increase in the frequency of the potentials. In all these respects the miniature potentials are like those of vertebrate muscles, except that, as Usherwood points out, the insect system seems to be rather less sensitive to magnesium ions. Since there seems to be much evidence for the statement that changes in frequency indicate presynaptic actions whereas changes in amplitude indicate postsynaptic actions (Katz, i962), an examination of the effects of calcium and magnesium ion concentration on the

16

D. J . AIDLEY

frequency of discharge of miniature potentials in Carausius might give some indication of the site of action of these ions in this case. Curare, acetylcholine and prostigmine had no effect on the miniature potentials, but 5hydroxytryptamine reduced their amplitude, resulting in complete block at high concentrations. Thus the pharmacology of

FIG. 5. Spontaneous miniature potentials from six different muscle fibres of Bluberus, recorded on a free-running trace. Calibrations 1 mV and 400 c/s. (From Usherwood, 1961.)

the miniature potentials appears to be similar to that of the neurally evoked postsynaptic potentials. After denervation of locust muscles, “ giant’’ miniature potentials up to 10 mV in height are observed at about the same time that impulse transmission from the nerve to the muscle fails (Usherwood, 1963b).

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It seems likely that these are connected with a disorganization of the motor nerve endings.

2. The quantal nature of the postsynaptic potential The presynaptic nerve endings at chemically transmitting synapses contain numbers of small vesicles. These are thought to contain “packets ” of transmitter substance which are released spontaneously in small numbers (producing miniature postsynaptic potentials) and in large numbers in the depolarization of the presynaptic nerve membrane. According to this interpretation, the postsynaptic potential is composed of a large number of quantal units, each resulting from the action of one of these packets on the postsynaptic membrane. The evidence for this is mainly derived from the observations of the sizes of the reduced postsynaptic potentials seen in frog muscles after treatment with Ringer solutions containing high magnesium ion concentrations. These potentials fluctuate in size by amounts equal to those of single miniature potentials. Similar “quantal ” fluctuations have been observed by Ushenvood (1963a,b) in postsynaptic potentials of locust muscle fibres after treatment with high magnesium ion concentrations or following denervation of the muscle. Presynaptic vesicles occur at insect neuromuscular junctions (Edwards et al., 1958). From these results, it seems that the intimate nature of the transmission process at insect neuromuscular junctions is similar to that at other chemically transmitting synapses. D . INHIBITORY RESPONSES

1. Peripheral inhibition in insects

Hoyle (1955b) showed that the locust metathoracic extensor tibiae muscle is innervated by three axons, one of which produces a small hyperpolarization of the muscle fibre membrane on stimulation. He was, however, unable to show that stimulation of this axon produced any inhibitory effect and suggested that its function was to enhance the amplitude of the “fast” electrical response so as to produce a larger contraction. As a consequence, the existence of peripheral inhibition in insects was until recently considered doubtful. Ripley and Ewer (1951) and Becht (1959) showed that the contraction of certain muscles could be reduced by increasing the intensity of the stimulus applied to the nerve trunk supplying the muscle, although in the absence of electrical records from the muscle, explanations other than inhibition are possible (see Hoyle, 1955b). More recently, Ikeda and

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D. J . AIDLEY

Boettiger (1965a,b) have demonstrated the presence of hyperpolarizing responses in the dorsoventral flight muscle of the bumble bee Bombus and in the basalar muscles of the rhinoceros beetle Oryctes, where it was shown that these hyperpolarizing potentials could cause attenuation of the excitatory postsynaptic potentials. The most complete study of this phenomenon is that performed by Usherwood and Grundfest (1964,1965) on the metathoracic extensor tibiae muscles of Schistocerca and Romalea, in which they were able to show both electrical and mechanical attenuation of the responses to excitatory stimuli; the rest of this section deals with their results. 2. Inhibitory postsynaptic potentials Stimulation of the inhibitory axon results in hyperpolarizing potentials with an amplitude of 1 to 25 mV (Fig. 6). These potentials summate on repetitive stimulation. In fibres in which the potentials are small in size, the potentials may show facilitation; in other cases this is not so.

IOOmsec

FIG.6. Intracellular records of the interaction of inhibitory and "slow" excitatory postsynaptic potentials in Schistocercu. The excitatory response was elicited at varying times after the inhibitory response. (From Usherwood and Grundfest, 1965.)

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By applying brief current pulses across the membrane via an intracellular electrode during the inhibitory postsynaptic potential, it was shown that the membrane conductance rises sharply at the beginning of the rising phase of the potential and then falls less rapidly. The duration of the inhibitory potentials is variable (40 to 300 msec.), but always longer than the excitatory potentials (15 to 150 msec.) in any one fibre. 3. Pharmacology of inhibition y-Aminobutyric acid (GABA) reduces the membrane resistance, causes hyperpolarization, and diminishes the excitatory postsynaptic potentials of fibres which are innervated by the inhibitor axon. Hence it apparently produces the same effects on the muscle fibre membrane as the inhibitory transmitter substance. Picrotoxin reverses these effects, and also diminishes the size of the inhibitory postsynaptic potentials. In these respects the pharmacology of inhibition in insects is identical to that of the crustacea (van der Kloot, 1960). 4. The ionic basis of inhibitory postsynaptic potentials

As mentioned earlier, the inhibitory potential is associated with an increase in membrane conductance. If the muscle membrane is hyperpolarized slightly, the inhibitory potential decreases in size, and further hyperpolarization results in the inhibitory potential being a depolarization. The level of membrane potential at which this changeover occurs (the reversal potential) is usually about - 70 mV (range - 55 to - 75 mV) and is thus near the value of the equilibrium potentials for potassium and chloride ions. In experiments in which all of the chloride in the saline solution was replaced by propionate, so altering the equilibrium potential for chloride ions, the inhibitory postsynaptic potentials reversed in sign. The reversal potential for the changes in membrane potential produced by GABA could be estimated by passing square current pulses through the membrane and finding at what potential the current-voltage curves obtained in the presence and absence of GABA crossed. This potential was -68 mV in potassium-free solutions and -67 mV in solutions with a potassium ion concentration of 30 mM, indicating that the reversal potential for the action of GABA was not the potassium equilibrium potential. It was concluded from these experiments that the action of the inhibitory transmitter substance, and of GABA (which may be the same thing), was to increase the permeability of the

20

D. J. AIDLEY

postsynaptic membrane to chloride ions. Here, again, the insect inhibitory system is similar to that of crustacea (Boistel and Fatt, 1958).

5. Interaction of inhibitory and excitatory responses Fig. 6 shows the effects of an inhibitory postsynaptic potential on a subsequent “slow” excitatory postsynaptic potential. Marked attenuation of the latter occurs if it arises during the initial phase of the inhibitory potential. Similar results have been obtained by Ikeda and Boetigger (1965b). This rather precise dependence of the interaction of the excitatory and inhibitory potentials on the timing of the two potentials may explain Hoyle’s failure to observe the phenomenon (Usherwood and Grundfest, 1964). The tetanic tension produced by stimulation of the slow axon in Romalea could be reduced by stimulation of the inhibitor axon, the attenuation increasing with increasing frequency of inhibitory stimulation. Complete inhibition of the mechanical response to stirhulation of the “slow” excitor axon at 12 impulses/sec. was obtained by stimulation of the inhibitor at 200/sec. In Schistocerca, such complete inhibition was never obtained, since not all of the muscle fibres supplied with the “slow” excitor axon are also innervated by the inhibitor.

Iv. THE ELECTRICAL EXCITABILITY OF THE MUSCLE FIBRE MEMBRANE A . T H E E L E C T R I C A L P R O P E R T I E S OF E L E C T R I C A L L Y E X C I T E D RESPONSES

When depolarizing current pulses are passed through the muscle fibre membrane by means of an intracellular electrode, the voltage recorded by a second electrode shows a further depolarization beyond that attributable to the resting resistance of the membrane jdel Castillo et a/., 1953; Cerf et a/., 1959). This electrically excited response differs from the propagated action potentials of nerve axons and vertebrate “twitch” muscle fibres in being a graded phenomenon (i.e. the size of the response is dependent upon the size of the stimulus). As a consequence, it is not propagated without decrement along the length of the fibre, but dies away within a few millimetres (Cerf et al., 1959). A similar phenomenon is seen in the responses to stimulation of the “fast” excitor axon in many preparations (del Castillo et al., 1953). In this case the stimulus for the production of the electrically excited response is the postsynaptic potential, and the point at which the

T H E E X C I T A T I O N OF I N S E C T S K E L E T A L MUSCLES

21

electrically excited response arises from the postsynaptic potential is given by the point of inflection on the rising phase of the response (Fig. 2). Here, again, the electrically excited component is a graded response, being proportional in size to the depolarization produced by the postsynaptic potential, as is evident in Fig. 4, where the size of the postsynaptic potential component is reduced in size by means of tryptamine. These electrically excited responses differ from the postsynaptic potentials in that they show refractoriness, so that there is a brief period after a response during which no second response can be elicited (the absolute refractory period) followed by a longer period during which the second response is reduced in size (the relative refractory period). Refractoriness occurs whether the electrically excited response is elicited by direct depolarization or by nervous stimulation (Cerf et al., 1959). The time course of recovery from refractoriness will probably be dependent on the size of the initial response, but this question remains to be investigated. If depolarizing pulses of long duration are passed through the membrane, the initial electrically excitable response is followed by a damped oscillation of the membrane potential (Cerf et al., 1959). B. T H E I O N I C B A S I S O F E L E C T R I C A L L Y E X C I T A B L E RESPONSES

The electrically excitable responses of insect muscle fibres are similar in type to the subthreshold “local responses” (Hodgkin, 1938) of nerve fibres. The well established theory of the ionic basis of the propagated action potential in squid giant axons (Hodgkin and Huxley, 1952) provides a convincing explanation of the mechanism of these responses. In a normal nerve action potential, depolarization causes an increase in the sodium conductance of the membrane which itself produces further depolarization, so that the membrane potential moves rapidly towards the sodium equilibrium potential. The return to the normal resting potential is brought about by two delayed consequences of depolarization: an increase in potassium conductance and a decrease in sodium conductance (the sodium inactivation process). Local responses arise because the increase in sodium conductance brought about by the subthreshold stimulus is too small and proceeds too slowly to counteract the effects of increase in potassium conductance. It seems very reasonable to suggest that the mechanism of the electrically excitable responses of insect muscle fibres is of the same type as the local responses of nerve axons (Hoyle, 1962). It is probable that the Hodgkin-Huxley equations would predict such responses if adjusted by

22

D . J . AIDLEY

suitable changes in the rate constants of the sodium and potassium activation processes. More specifically, it has been suggested that the increase in potassium conductanceis more rapid in the graded responses of insect muscles (Werman et ul., 1961). The presence of a sodium inactivation process operative at the normal resting potential seems to be implied by some results of Cerf et ul. (1959), in which the electrically excitable response arose at a much more negative membrane potential after the fibre had been hyperpolarized for a short time. While the general principles of this argument seem to be fairly sound, it is not so clear that the ion involved in the specific, regenerative increase in permeability (i.e. the ion responsible for the rising phase of the electrically excitable response) is sodium, or sodium alone. Wood (1963) has investigated the effects of sodium ion concentration in the size of the electrical responses to “fast” axon stimulation in the leg muscles of Locustu, Periplunetu and Curuusius, at the same time measuring the internal sodium concentrations of muscles subjected to similar treatments, so that estimates of the sodium equilibrium potential could be made. In Locustu, the peak of the response occurred at an average membrane potential of - 19 mV in a solution containing no sodium ions, rising to + 5 mV in a solution with a sodium ion concentration of 150 m ~The . change was rather less for PeripZunetu (the corresponding figures were - 15 and + 2 mv), and much less for Caruusius (-3 and +4.5 mv). The values obtained at nominally zero sodium ion concentrations are interesting in that they are more positive than the sodium equilibrium potential, even if the actual sodium ion concentration were as much as 5 m~ (which is most unlikely). Interpretation of these results is complicated by the fact that the quantity measured was total electrical responses to nervous stimulation, which would include (and in zero sodium salines may entirely consist of) a large postsynaptic potential component. Since sodium is the major cation in the haemolymph of Locustu and PeripZunetu and since the membrane potential at the peak of the response is moderately sensitive to the sodium ion concentration, it is probable that sodium is the major ion involved in the rising phase of the electrically excitable response, although participation of other cations, such as calcium, is not excluded. In Curuusius, however, where the haemolymph sodium concentration is only 15 m~ (giving a sodium equilibrium potential of about +4.6 mV; Wood, 1963) and the potential at the peak of the response is only slightly affected by the sodium ion concentration, it seems rather unlikely that sodium is the only, or indeed the major ion involved. In view of the dependence of the amplitude of the electrical response to stimulation

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on the magnesium ion concentration, and the very high magnesium ion concentration in the haemolymph, it seems probable that magnesium carries the action current in this case (Wood, 1958). The observations of Treherne (1965) on nervous conduction in the abdominal nerve cord of Carausius are of interest in this connection; he found that nervous conduction ceased in the absence of either sodium or magnesium ions. If the conclusion that magnesium ions carry a major part of the action current in Carausius is correct, then there is a very high probability that an active transport system concerned with maintaining the magnesium ion concentration gradient across the cell membrane exists. C . SPONTANEOUS ACTIVITY

If locust muscle fibres are perfused with calcium- and magnesium-free saline solutions, there follows a brief period during which the membrane is hyperexcitable, producing repetitive spike-like depolarizations (D. J. Aidley, unpublished). Similar effects have been seen in frog skeletal muscle (Bulbring et al., 1956). Van der Moot (1963) has reported some interesting observations on the spiracular muscles of the pupae of the silkmoths Hyalophora and Telea. These undergo spontaneous activity which consists of slow depolarizations of the membrane (pacemaker potentials), each followed by a spike-like component. The pacemaker potentials were accompanied by an increase in membrane conductance. If the membrane potential was set to various levels by passing current through an intracellular electrode, the pacemaker potentials always reached a plateau at the same value, - 29 mV. This suggests that the potentials are produced by an increase in the membrane conductance for some ion which has an equilibrium potential at - 29 mV, but what this ion is is far from clear. If the membrane is depolarized beyond - 29 mV, the potentials reverse in sign; this is rather extraordinary, since it implies that the cyclic nature of the phenomenon is based on changes in membrane permeability which are not determined by changes in membrane potential. V. THE EXCITATION-CONTRACTION COUPLING PROCESS A . DEPOLARIZATION

The evidence that depolarization of the cell membrane is necessary for the contraction of the skeletal muscles of vertebrates (Kuffler, 1946; Sten-Knudsen, 1960) and crustacea (Orkand, 1962) is considerable. The same situation appears to exist in insect skeletal muscles. 2-kA.I.P.

4

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D. J. A I D L E Y

If the amplitude of the electrical response to nervous stimulation is reduced by means of neuromuscular blocking agents, the twitch tension is also reduced, as is shown in Fig. 4. Perfusion of a muscle with salines containing high potassium ion concentrations causes depolarization and contraction. In the metathoracic spiracular muscle of locusts (Hoyle, 1961), the threshold for the development of this contracture occurs at a potassium ion concentration of about 30 m~ (membrane potential - 34 mv) ; contracture tension increases with increasing potassium ion concentration to reach the maximum level at about 70 m~ (membrane potential - 18 mv). A similar result has been obtained from the mesothoracic extensor tibiae of Schistocerca (Aidley, 1965a). A more subtle influence of membrane potential on tension development is seen in frog “ twitch ” fibres, where the potassium contracture lasts only for a few seconds (Hodgkin and Horowicz, 1960). If the membrane is then repolarized by perfusion with a solution containing a lower potassium ion concentration, a restorative (“priming”) process occurs, so that the fibre is again able to contract on depolarization. The extent and rapidity of this “priming” process increases with increasing (more negative) membrane potentials. Potassium contractures of a similar short duration occur in the metathoracic extensor tibiae (Hoyle, 1961) and flexor tibiae (D. J. Aidley, unpublished) of Schistocerca, and it is not unlikely that a “priming” process similar to that described by Hodgkin and Horowicz could be demonstrated in these and some other insect muscles. B. THE I M P O R T A N C E O F C A L C I U M IONS

If the mesothoracic extensor tibiae of Schistocerca is perfused with a calcium-free solution containing the chelating agent ethylenediaminetetra-acetate (EDTA), it will no longer contract when depolarized by an isotonic solution of potassium chloride (Aidley, 1963, 1965a). In this condition, contraction can be elicited by addition of calcium chloride to the potassium chloride solution bathing the muscle (Fig. 7). The effect of solutions containing various concentrations of calcium ions (in the absence of EDTA) is slower, but it can be shown that the maximum contracture tension is related to the external calcium ion concentration, with a threshold at about 0.03 m~ and a saturation value in the region 2 to 4 mM. Thus it seems that insect muscles are similar to vertebrate muscles (Frank, 1960; Niedergerke, 1956; Edman and Schild, 1962) in requiring calcium ions for contraction. In vertebrate muscles, the presence of calcium ions in the sarcoplasm appears to be necessary for the interaction of actin, myosin and adeno-

-

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25

sine triphosphate so as to produce contraction, and the calcium ion concentration is probably controlled by a “relaxing factor” system composed of vesicular elements derived from the endoplasmic reticulum (see, for example, Needham, 1960, and Weber et al., 1964). A relaxing factor whose action is antagonized by calcium ions has been isolated from locust skeletal muscles by Tsukamoto and his colleagues (quoted in Maruyama, 1965). The contraction of glycerinated fibres from the

A -

C

-

7 ]log

5 min

FIG.7. The effects of treatment with EDTA and subsequent addition of calcium chloride on the potassium contracture of the mesothoracic extensor tibiae of Schistocerca. The horizontal line under each record shows the duration of perfusion with KCl solution. Before each perfusion with KCl solution the muscle was treated as follows: (A) perfusion with a standard Ringer solution (4 mM CaCI2); (B) perfusion for 40 min. with a Ca-free Ringer containing 4 m~ EDTA; and, (C) perfusion for a further 60 min. with standard Ringer. The plateau of a tetanus produced by stimulation of the motor nerves is shown at the beginning of record (A). (From Aidley, 1965.)

fight muscles of the giant water bugs Lethoceros and Hydrocirius in the presence of adenosine triphosphate is dependent upon the presence of calcium ions (Jewell et al., 1964). This contractile action of calcium ions appears to be rather specific, in that strontium and a number of other divalent cations do not form adequate substitutes for calcium in the promotion of potassium contracture (Aidley, 1965b). Application of solutions containing strontium ions to calcium-depleted muscles depolarized by potassium chloride solution results in submaximal, phasic contractures;but successive$responses to depolarization decline in the continued absence of calcium

26

D . J . AIDLEY

ions, suggesting that the small contractions mediated by strontium ions are brought about by a displacement of calcium ions from some intracellular store. Similar results have been obtained by Frank (1962) on frog toe muscles. C . THE ACTION OF CARBON DIOXIDE

Hoyle (1961) has shown that application of carbon dioxide to the metathoracic spiracular muscle of Schistocerca causes relaxation when the muscle is in potassium contracture, although the membrane potential of the depolarized fibres was unaffected. This result implies that carbon dioxide has some direct effect on the coupling process in this muscle in addition to its effects on the resting membrane and the size of the postsynaptic potential. Contracture tension of the spiracular muscle is maintained for many hours in isotonic solutions of potassium chloride or sulphate, but falls to zero in about half an hour in isotonic potassium bicarbonate (Hoyle, 196 1). The mechanisms of the relaxations occurring in the presence of bicarbonate or carbon dioxide are not known, although it is conceivable that bicarbonate acts by reducing the calcium ion concentration. Further evidence for a direct action of carbon dioxide on the coupling process of the locust spiracular muscle has been adduced by Hoyle from experiments in which he measured the effects of a brief “pulse” of carbon dioxide on the size of the postsynaptic potentials of individual fibres and the twitch tension of the whole muscle (Hoyle, 1960). It was found that a “hysteresis” effect occurred, in which the size of the postsynaptic potential fell and rose more rapidly than the twitch tension. These experiments cannot be considered conclusive, however, since the whole cycle of depression and recovery lasted only a few seconds, a time which is probably comparable with the diffusioii delay which must occur between the carbon dioxide affecting fibres in different parts of the muscle. D.

“

FAST

”

AND

“

SLOW

”

MUSCLES

Skeletal muscle fibres of frogs can be divided into two clearcut types, known as “twitch” (or “fast”) and “slow” fibres. Some muscles, such as the sartorius, are composed entirely of “fast” fibres, others, such as the iliofibularis and rectus abdominis, contain a high proportion of “slow” fibres (Kuffler and Vaughan Williams, 1953). “Slow” fibres differ from “fast” fibres in that they are multiterminally innervated, electrically inexcitable, contract at much slower velocities and do not

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relax after a few seconds when immersed in solutions containing high potassium ion concentrations. These distinctions cannot be usefully applied to insect muscles, which in many cases can be classified as either “fast” or “slow” according to the criteria used. For example, if we compare the metathoracic and mesothoracic extensor tibiae muscles of locusts, we find that the fibres of both muscles are electrically excitable (as in “fast” muscles), that they are both multiterminally innervated (as in “slow” muscles), and that the potassium contracture is brief (as in “fast” muscles) in the metathoracic muscle whereas it is maintained (as in “slow” muscles) in the mesothoracic muscle (Hoyle, 1961; Aidley, 1963). Becht and Dresden (1956) suggested that the coxal muscles of the cockroach could be divided into two types (termed “fast” and “slow”) on the basis of their mechanical responses to stimulation. However, later investigations using more sophisticated techniques failed to uphold this view (Becht et al., 1960; Usherwood, 1962). It is clear that there are differences in the mechanical properties of various insect muscles, but a clearcut division into two types is not at present possible. I am very grateful to Dr. P. N. R. Usherwood and Dr. A. C. Neville for their comments on an early draft of this article. REFERENCES

Adrian, R.H. (1956). The effect of internal and external potassium concentration on the membrane potential of frog muscle. J. Physiol. 133, 631-658. Aidley, D. J. (1963). Influence of calcium ions on potassium contracture in an insect leg muscle. Nature, Lond. 198,591-592. Aidley, D.J. (1965a). The effect of calcium ions on potassium contracture in a locust leg muscle. J. Physiol. 177, 94-102. Aidley, D.J. (1965b). The effects of strontium and other divalent cations on potassium contracture in a locust leg muscle. J. Physiol. 177, 103-111. Becht, G. (1959). Studies on insect muscles. Bijdr. Dierk. 29, 5-40. Becht, G. and Dresden, D. (1956). Physiology of the locomotory muscles in the cockroach. Nature, Lond. 177,836-837. Becht, G., Hoyle, G. and Usherwood, P. N. R. (1960). Neuromuscular transmission in the coxal muscles of the cockroach. J. Insect Physiol. 4, 191-201. Belton, P. (1958). Membrane potentials recorded from moth muscle fibres. J. Physiol. 142,20P. Belton, P. (1960). Effect of ions on potential in lepidopteran muscle fibres. Biol. Bull. mar. biol. Lab., Woods Hole 119, 289. Belton, P.and Grundfest, H. (1962). Potassium activation and K spikes in muscle fibres of the mealworm larva (Tenebrio molitor). Am. J. Physiol. 203, 588594.

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Boistel, J. and Fatt, P. (1958). Membrane permeability change during transmitter action in crustacean muscle. J. Physiol. 144, 176-191. Boyle, P. J. and Conway, E. J. (1941). Potassium accumulation in muscle and associated changes. J. Physiol. 100, 1-63. Bulbring, E.,Holman, M. and Lullman, H. (1956). Effects of calcium deficiency on striated muscle of the frog. J. Physiol. 133, 101-1 17. Cerf, J. A., Grundfest, H., Hoyle, G. and McCann, F. V. (1959). The mechanism of dual responsiveness in muscle fibres of the grasshopper Romalea microptera. J . gen. Physiol. 43, 377-395. Conway, E. J. (1957). Nature and significance of concentration relations of potassium and sodium ions in skeletal muscle. Physiol. Rev. 37, 84-132. del Castillo, J. and Engbaek, L. (1954). The nature of the neuromuscular block produced by magnesium. J. Physiol. 124, 370-384. del Castillo, J. Hoyle, G. and Machne, X. (1953). Neuromuscular transmission in a locust. J. Physiol. 121, 539-547. del Castillo, J. and Katz, B. (1954). Quanta1 components of the end-plate potential. J. Physiol. 124, 560-573. Duchgteau, G., Florkin, M. and Leclercq, J. (1953). Concentrations des bases fixes et des types de compositions de la base totale de l’hemolymph des insectes. Arch. int. Physiol. 61, 518-549. Dudel, J. and Kuffler, S. W. (1961). Mechanism of facilitation at the crayfish neuromuscular junction. J. Physiol. 155,530-542. Edman, K. A. P. and Schild, H 0. (1962). The need for calcium in the contractile responses induced by ecetylcholine and potassium in the rat uterus. J. Physiol. 161,4244ll. Edwards, G. A., Ruska, H. and de Harven, E. (1958). Neuromuscular junctions in flight and tymbal muscles of the Cicada. J. biophys. biochem. Cytol. 4,251-255. Fatt, P. and Katz, B. (1951). An analysis of the end-plate potential recorded with an intracellular electrode. J. Physiol. 115,320-370. Fatt, P. and Katz, B. (1952). Spontaneous subthreshold activity at motor nerve endings. J. Physiol. 117, 109-128. Fatt, P. and Katz, B. (1953). The electrical properties of crustacean muscle fibres. J. Physiol. 120, 171-204. Frank, G. B. (1960). Effects of changes in extracellular calcium concentration on the potassium-induced contracture of frog’s skeletal muscle. J. Physiol. 151, 518-538. Frank, G. B. (1962). Utilization of bound calcium in the action of caffeine and certain multivalent cations on skeletal muscle. J. Physiol. 163,254-268. Hagiwara, S. and Watanabe, A. (1954). Action potential of insect muscle examined with intracellular electrode. Jap. J. Physiol. 4, 65-78. Harlow, P. A. (1958). The action of drugs on the nervous system of the locust (Locusta migratoria). Ann. appl. Biol. 46,55-73. Hill, R.B. (1963). The effects of acetylcholine on twitches in the locust leg. Comp. Biochem. Physiol. 10, 203-208. Hill, R. B. and Usherwood, P. N. R. (1961). The action of 5-hydroxytryptamine and related compounds on neuromuscular transmission in the locust, Schistocerca gregaria. J. Physiol. 153,393-401. Hodgkin, A. L. (1938). The subthreshold potentials in a crustacean nerve fibre. Proc. R. SOC.B 126,87-121.

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Hodgkin, A. L. (1951). The ionic basis of electrical activity in nerve and muscle. Biol. Rev. 26, 339-409. Hodgkin, A. L. and Horowicz, P. (1959). The influence of potassium and chloride ions on the membrane potential of single muscle fibres. J. Physiol. 148, 127-160. Hodgkin, A. L. and Horowicz, P. (1960). Potassium contractures in single muscle fibres. J. Physiol. 153,386-403. Hodgkin, A. L. and Huxley, A. F. (1952). A quantitative description of conduction and excitation in nerve. J. Physiol. 117, 500-544. Hodgkin, A. L. and Katz, B. (1949). The effect of sodium ions on the electrical activity of the giant axon of the squid. J. Physiol. 108,37-77. Hoyle, G.(1953). Potassium ions and insect nerve muscle. J. exp. Biol. 30,121-135. Hoyle, G . (1955a). The anatomy and innervation of locust skeletal muscle. Proc. R. SOC.B 143,281-292. Hoyle, G. (1955b). Neuromuscular mechanisms of a locust skeletal muscle. Proc. R. SOC.B 143,343-367. Hoyle, G. (195%). The effects of some common cations on neuromuscular transmission in insects. J. Physiol. 127, 90-103. Hoyle, G. (1957). Nervous control of insect muscles. In “Recent Advances in Invertebrate PhysiQlogy ” (B. T. Scheer, ed.), pp. 73-98. University of Oregon Publications. Hoyle, G. (1 959). The neuromuscular mechanism of an insect spiracular muscle. J. Insect Physiol. 3, 378-394. Hoyle, G.(1960). The action of carbon dioxide gas on an insect spiracular muscle. J. Insect Physiol. 4, 63-79. Hoyle, G.(1961). Functional contracture in a spiracular muscle. J. Insect Physiol. 7, 305-314. Hoyle, G . (1962). Comparative physiology of conduction in nerve and muscle. Am. Zoologist 2, 5-25. Hoyle, G. (1965). Neural control of ske!etal muscle. In “The Physiology of Insects" (M. Rockstein, ed.) vol. 2, pp. 407-449. Academic Press, New York and London. Ikeda, K. and Boettiger, E. G. (1965a). Studies on the flight mechanism of insects. 11. The innervation and electrical activity of the fibrillar muscles of the bumble bee, Bombus. J. Insect Physiol. 11, 779-789. Ikeda, K. and Boettiger, E. G. (1965b). Studies on the flight mechanism of insects. 111. The innervation and electrical activity of the basalar fibrillar flight muscle of the beetle, Oryctes rhinoceros. J. Insect Physiol. 11, 791-802. Jenden, D. J. and Reger, J. F. (1963). The role of resting potential changes in the contractile failure of frog sartorius muscles during calcium deprivation. J. Physiol. 169,889-901. Jewell, B. R., Pringle, J. W. S. and Riiegg, J. C. (1964). Oscillatory contraction of insect fibrillar muscle after glycerol extraction. J. Physiol. 173,6-8P. Katz, B. (1962). The transmission of impulses from nerve to muscle, and the subcellular unit of synaptic action. Proc. R. SOC.B 155, 455-479. Kerkut, G. A. and Ridge, R. M. A. P. (1961). The effect of temperature changes on the resting potential of crab, insect and frog muscle. Comp. Biochern. Physiol. 3, 64-70.

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Kerkut, G. A., Leake, L. D., Shapira, A., Cowan, S. and Walker, R. J. (1965b). The presence of glutamate in nerve-muscle perfusates of Helix, Carcinus, and Periplaneta. Comp. Biochem. Physiol. 15,485-502. Kerkut, G. A., Shapira, A. and Walker, R. J. (1965a). The effect of acetylcholine, glutamic acid and GABA on the contractions of the perfused cockroach leg. Comp. Biochem. Physiol. 16, 37-48. Keynes, R. D. (1963). Chloride in the squid giant axon. J. Physiol. 127, 90-103. Kuffler, S. W. (1946). The relation of electrical potential changes to contracture in skeletal muscle. J. Neurophysiol. 9, 367-377. Kuffler, S. W. and Vaughan Williams, E. M. (1953). Properties of the “slow” skeletal muscle fibres of the frog. J. Physiol. 121, 318-340. Maruyama, K. (1956). The biochemistry of the contractile elements of insect muscle. In “The Physiology of Tnsecta” (M. Rockstein ed.) Vol 2, pp. 451-483. Academic Press, New York and London. McCann, F. V. and Boettiger, E. G. (1961). Studies on the flight mechanism of insects. I. The electrophysiology of fibrillar flight muscle. J. gen. Physiol. 45, 1 25-1 42. Needham, D. M. (1960). Biochemistry of muscular action. In “The structure and function of muscle” (G. H. Bourne, ed.) Vol. 2, pp. 55-104. Academic Press, London and New York. Neville, A. C. (1963). Motor unit distribution of the dorsal longitudinal flight muscles in locusts. J. exp. Biol. 40, 123-136. Niedergerke, R. (1956). The potassium chloride contracture of the heart and its modification by calcium. J. Physiol. 134, 584-599. Orkand, R. K. (1962). The relation between membrane potential and contraction in single crayfish muscle fibres. J. Physiol. 161, 143-159. Pringle, J. W. S. (1939). The motor mechanism of the insect leg. J. exp. Biol. 16,220-23 1. Ripley, S. H. and Ewer, D. W. (1951). Peripheral inhibition in the locust. Ncture, Lond. 167, 1066. Sten-Knudsen, 0. (1960). Is muscle contraction initiated by internal current flow? J. Physiol. 151, 363-384. Takeuchi, A. and Takeuchi, N. (1960). On the permeability of the end-plate membrane during the action of transmitter. J. Physiol. 154, 52-67. Treherne, J. E. (1965). Some preliminary observations on the effects of cations on conduction processes in the abdominal nerve cord of the stick insect, Carausius morosus. J. exp. Biol. 42, 1-6. Usherwood, P. N. R. (1961). Spontaneous miniature potentials from insect muscle fibres. Nature, Lond. 191, 814-815. Usherwood, P. N. R. (1962). The nature of “slow” and “fast “ contractions in the coxal muscles of the cockroach. J. Insect Physiol. 8, 31-52. Usherwood, P. N. R. (1963a). Spontaneous miniature potentials from insect muscle fibres. J. Physiol. 169, 149-160. Usherwood, P. N. R. (1963b). Response of insect muscle to denervation. 11. Changes in neuromuscular transmission. J. Insect Physiol. 9, 81 1-125. Usherwood, P. N. R. and Grundfest, H. (1964). Inhibitory postsynaptic potentials in grasshopper muscle. Science, N. Y. 143, 817-818 Usherwood, P. N. R. and Grundfest, H. (1965). Peripheral inhibition in skeletal muscle of insects. J. Neurophysiol. 28, 497-518.

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van der Kloot, W. G. (1960). Picrotoxin and the inhibitory system of crayfish muscle. In “Inhibition in the nervous system and y-aminobutyric acid” (E. Roberts, ed.) pp. 409-412. Pergamon Press, New York and Oxford. van der Kloot, W. G. (1963). The electrophysiology and the nervous control of the spiracular muscle of pupae of giant silkmoths. Comp. Biochem. Physiol. 9,317-333.

Weber, A., Herz, R. and Reiss, I. (1964). The regulation of myofibrillar activity by calcium. Proc. R. SOC.B 160, 489-499. Werman, R., McCann, F. V. and Grundfest, H. (1961). Graded and all-or-none electrogenesis in arthropod muscle. I. The effects of alkali-earth cations on the neuromuscular system of Romalea microptera. J. gen. Physiol. 44, 979-995.

Wood, D. W. (1957). The effect of ions upon neuromuscular transmission in a herbivorous insect. J. Physiol. 138, 119-139. Wood, D. W. (1958). The electrical and mechanical responses of the prothoracic flexor tibialis muscle of the stick insect. J. exp. Biol. 35, 850-861. Wood, D. W. (1961). The effect of sodium ions on the resting and action potentials of locust and cockroach muscle fibres. Comp. Biochem. Physiol. 4, 42-46.

Wood, D. W. (1963). The sodium and potassium composition of some insect skeletal muscle fibres in relation to their membrane potentials. Comp. Biochem. Physiol. 9, 151-159. Wood, D. W. (1965). The relationship between chloride ions and resting potential in skeletal muscle fibres of the locust and the cockroach. Comp. Biochem. Physiol. 15, 303-312.

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The Excretion of Nitrogen in Insects E. BURSELL Department of Biological Sciences, University College of Rhodesia and Nyasaland, Salisbury, Rhodesia I. Introduction . 11. The Formation of Nitrogenous End Products A. The uricolytic pathway . B. The uricotelic pathway. . C. The formation of urea D. The formation of ammonia . E. Amino acids . F. Miscellaneous materials . LU. The Excretion of Nitrogenous End Products A. Collembola . B. Orthoptera . C. Odonata . D. Dermaptera . E. Hemiptera F. Coleoptera G. Neuroptera . H. Hymenoptera . I. Diptera . J. Lepidoptera . N. Conclusions . References .

.

. .

33 34 36

.

41 42 43

. 4 0

. .

. 4 4 .

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

4

4 45 46 47 48 48 50 51 51 52 54 56 61

I. INTRODUCTION The occurrence of uric acid in the excreta of insects was established before the turn of the century and its presence has been confirmed subsequently by a large number of investigators (see reviews in Wigglesworth, 1950; Prosser, 1952; Roeder, 1953; Chauvin, 1956). Earlier findings were generalized in statements such as those of Needham (1935): “. . . in the insect, excretion of uric acid as the main end product of nitrogen metabolism is widespread, if not universal”, and of Florkin (1945): “I1 est bien Ctabli que chez les insectes adultes, l’acide urique reprksente le terme pridominant du catabolisme protidique”. On this basis the insect was assessed as being predominantly uricotelic in a general classification of excretory metabolism (Needham, 1950). Uricotelism was seen as an adaptation to a terrestrial mode of life; the 33

34

E. B U R S E L L

excretion of toxic end products, like ammonia, or of soluble end products, like urea, was seen as being militated against by the shortage of water. This view of the insects as a class still prevails (e.g. reviews by Craig, 1960; Stobbart and Shaw, 1964), subject to certain exceptions. In recent years, however, data have accumulated which suggest that nitrogenous end products other than uric acid may figure largely in the excreta of a variety of insects; the exceptions are becoming more numerous, and the validity of the earlier generalization appears to be threatened. In the following account, an attempt has been made to summarize the available information, and, in this way, to provide some basis for a reassessment of the situation. 11. THE FORMATION OF NITROGENOUS END P R O D U C T S The metabolic interrelationships of the main nitrogenous end products in insects, and the sources from which they derive, are summarized in Fig. 1. Two essentially opposing metabolic pathways appear to be involved.

(a) One has its origin in the nucleic acids, and may be termed the uricolytic pathway. It involves deamination and oxidation of the purine raw material (adenine and guanine) followed by the breakdown of the ring system to progressively simpler nitrogenous waste products. The last steps are thought to play little, if any, part in the excretory metabolism of insects, and they have been indicated by dotted lines. (b) The other has its origin in the proteins and their constituent amino acids, and may be termed the uricotelic pathway. It involves the synthesis of purines from amino acid nitrogen. In some insects the uricotelic pathway appears to be inoperative, and amino acid nitrogen is excreted as ammonia. In many others, urea appears as a minor constituent of the excreta, being apparently derived from protein nitrogen. As a third possibility, in certain insects specific amino acids are excreted as such; these alternative pathways are indicated in the diagram. Values for the nitrogen content and the solubility of nitrogenous end products have been included in Fig. 1. Ammonia is highly soluble, but a most efficient vehicle for the removal of waste nitrogen, with a nitrogen content of more than 80%; urea contains substantially less nitrogen and is of the same order of solubility as ammonia. The purines and their near derivatives contain between 30 and 40% nitrogen, but are

35

THE EXCRETION O F NITROGEN I N INSECTS

very much less soluble than ammonia and urea, while two of the amino acids listed have a nitrogen content not much lower than the purines, but are relatively soluble. Further reference to these differences and

I Nucleic Acid I

260

33

6

Allantoin

35

60

Allantoic Acid

32

slightly soluble

46

119,300

82

89,000

32 27 6

15,000 soluble

Uric Acid

’’ Arginine Histidine Cystine

Solubility mg/lOO ml

37

Xanthine

I Protein HAmino Acids I

% Nitrogen

Ammonia

.

FIG. 1. Nitrogenous end products and their metabolic interrelationships. (Data from “Handbook of Chemistry and Physics” ed. Hodgman, C. D., Chemical Rubber Publishing Co., Cleveland, Ohio.)

their implications in relation to nitrogenous excretion will be deferred until the excretory metabolism of different insects has been reviewed. The pyrimidines constitute another nitrogenous component of nucleic acid, but little information is available concerning their metabolism in insects. Recent studies (Bruno and Cochran, 1965) suggest

11

36

E. B U R S E L L

that it may conform in part to the vertebrate pattern, but further work will need to be done before their possible role as nitrogenous end products can be assessed. A . THE URICOLYTIC P A T H W A Y

The starting material for this reaction chain is nucleic acid, arising in part as a product of digestion, and in part from processes of tissue

Adenine

Hypoxanthine

1

(xanthine dehyhogenase)

Xanthine

Guanine

1

(xanthine dehydrogenase)

I

Uric Acid

maintenance and repair. The primary breakdown products are the nucleosides adenosine and guanosine, and these may be deaminated directly under the influence of the corresponding enzymes, adenosine deaminase and guanosine deaminase. In the presence of purine nucleosidases, the ribose sugar is then split off to liberate the purine bases.

T H E EXCRETION OF N I T R O G E N I N INSECTS

37

Enzymes responsible for such actions have been demonstrated in the cockroach (Cochran and Bruno, 1963; Cordero and Ludwig, 1963), and adenosine deaminase activity has been established in extracts of the larval fat body of blowflies (Desai and Kilby, 1958a) and in homogenates of Drosophila (Wagner and Mitchell, 1948). On the other hand, Duchlteau et al. (1940) were unable to show the presence of adenosine or guanosine deaminase in certain insects, and it is possible that in such species deamination occurs after hydrolysis of the nucleosides, as shown opposite. Adenase and guanase, the enzymes responsible for deamination of the free purine bases, have been shown to occur in a number of different insects (Duchfiteau et al., 1940; Desai and Kilby, 1958a; Prota, 1961). In extracts from the fat body of the silkworm the activity of guanase is much greater than that of adenase (Hayashi, 1961), and this appears to be so in other insects as well (Anderson and Patton, 1955). The deaminated purines, xanthine and hypoxanthine, are oxidized to uric acid under the influence of a single enzyme, xanthine oxidase, or xanthine dehydrogenase as it should more properly be called in view of the investigations of Irzykiewicz (1955); this author has shown that, unlike its mammalian counterpart, the insect enzyme cannot use molecular oxygen, but can reduce methylene blue. It is possible that nicotinamide-adenine dinucleotide (NAD) may act as the hydrogen acceptor for this reaction under normal conditions. Early workers failed to find evidence of the oxidation of xanthine in insects (Truszkowski and Chajkinowna, 1935), but xanthine dehydrogenase activity has subsequently been demonstrated in a variety of species, and the enzyme appears to be widely distributed throughout the class (Duchlteau et al., 1940; Florkin and Duchlteau, 1941; Anderson and Patton, 1954; Desai and Kilby, 1958b; Lisa and Ludwig, 1959; Ursprung and Hadorn, 1961; Hayashi, 1962; Cordero and Ludwig, 1963; Keller and Glassman, 1963; Parzen and Fox, 1964). In fact, there are indications that there may be more than one molecular form of the enzyme in Drosophila (Keller et al., 1963; Smith et al., 1963). The uric acid formed under the influence of xanthine dehydrogenase may be subjected to a series of degradative reactions in the presence of uricolytic enzymes as summarized overleaf. The first step is the oxidation of uric acid to allantoin under the iduence of the enzyme uricase. This enzyme has long been known to be active in insects (Leifert, 1935; Truskowski and Chajkinowna, 1935;Brown, 1938a; ROCCO, 1938),and more recent work has confirmed

38

E. BURSELL

its wide distribution within the class (Razet, 1952,1953, 1954,1956,1957, 1961;Lisa and Ludwig, 1959; Corder0 and Ludwig, 1963; Nelson, 1964), though it has been reported absent in some species (Florkin and DuchC teau, 1943; Razet, 1961). In Popillia japonica uricase is present in extracts of the embryo and of larval stages, but absent from the prepupa and pupa (Ross, 1959); a similar loss of uricolytic activity during pupal development has been reported for the blowfly Lucilia (Brown, 1938a).

I I

Allantoic acid O=C

(uricase)

(allantoinme)

YH2 COOH I \,cH. H

N\H2

c=o

N’

H

1

(allanroicme)

COOH

I

CHO

/NH2

-t-2O=C,

Glyoxylic acid

NH, Urea

I

I

(weme)

2 C 0 2 + 4NHs

T H E E X C R E T I O N OF N I T R O G E N I N I N S E C T S

39

The hydrolysis of allantoin to allantoic acid, which constitutesa further breakdown of the purine ring system, is catalysed by the enzyme allantoinase. Active extracts of this enzyme have also been obtained from a variety of insects (Rocco, 1936 and 1938; Manunta, 1948 and 1949; Razet, 1952-1957), but it appears to be somewhat less widespread than uricase (Razet, 1961). The next step in the sequence of uricolytic breakdown is the hydrolysis of allantoic acid under the influence of allantoicase, leading to the formation of urea and glyoxylic acid. This enzyme appears to have a very restricted distribution. It has been reported for larval stages of Bombyx (Manunta, 1948), but Razet (1961) failed to confirm its presence in this species. Feebly active preparations have been made from insects belonging to a number of different orders, but only from two of the many species tested by Razet (1961) have highly active extracts been obtained, namely from Xenylla wekhii (Collembola) and Achetu domesticu (Orthoptera). Since urea is absent from the excreta, or present in very small amounts, even in those insects which possess an active allantoicase, it seems safe to conclude that uricolytic breakdown beyond the stage of allantoicacid is of little significancein the excretorymetabolism of insects. The last in the chain of uricolytic enzymes is urease, which catalyses the hydrolysis of urea to ammonia and carbon dioxide. The presence of this enzyme has not been unequivocally established in insects. It has been reported to occur in the meat-eating larvae of certain Diptera (Tomita and Kumon, 1936; Baker, 1939; Robinson and Baker, 1939; Robinson and Wilson, 1939), and in a small number of other insects (Razet, 1961), but the methods used for its detection have not been above reproach, and the evidenceshould be accepted with caution. Lennox (1941) failed to find trace of its existence in a careful study of the enzymes responsible for ammonia production in larvae of the blowfly Luciliu cuprina. Even if the presence of urease were accepted for the species concerned, it is doubtful if it could be considered as a uricolytic enzyme, since earlier members of the chain are missing (Razet, 1961). A considerable amount of work has been done on the localization of enzymes involved in purine metabolism. The activity of xanthine dehydrogenase is high in the fat body of Peripluneta (Anderson and Patton, 1954 and 1955), and in larvae of Drosophila the fat body is more active than the Malpighian tubules (Ursprung and Hadorn, 1961). In Tenebrio the enzyme appears to be more concentrated in extracts from the mid gut and fat body than from other tissues (Irzykiewicz, 1955). Uricase has also been shown to occur in the fat body ofinsects (Leifert, 1935; Lisa and Ludwig, 1959; Pierre, 1964), but in the species studied by Razet

40

E. BURSELL

(1961) highest activities were usually found in extracts of the Malpighian tubules, with substantially lower concentrations occurring in mid gut and fat body extracts; and in Musca domestica, too, uricase activity is concentrated in the Malpighian tubules (Nelson, 1964). In insects which possess an active allantoinase, the enzyme distribution appears to be similar to that of uricase, with highest activity in the Malpighian tubules, followed by fat body and mid gut (Razet, 1961). B . T H E URICOTELIC P A T H W A Y

While the breakdown of uric acid in insects has been the subject of extensive investigations during the last decade, its synthesis has received relatively little attention. It has long been realized that the large amounts of uric acid which occur in the excreta of insects cannot derive solely from nucleic acid sources; the bulk of it must originate from protein nitrogen, but the synthetic pathways involved have not yet been fully elucidated. An early suggestion was made that arginine and histidine might be involved as intermediaries in uric acid synthesis (see Baldwin, 1948; Hoskins and Craig, 1935),but the existence of such a pathway has not been confirmed. The possibility of identifying precursors of uric acid in insects has been investigated both in vivo (Brighenti and Colla, 1940) and in vitro (Leifert, 1935;Anderson and Patton, 1955; Desai and Kilby, 1958b), and various non-purine substances including urea, malonate, ammonium salts, monoethyloxaloacetate and 4-amino-5-imidazole carboxamide riboside have been reported to stimulate the production of uric acid in certain cases. A more direct approach to the problem was made by McEnroe and Forgash (1957 and 1958) who investigated the fate of radioactive formate in Periplaneta umericana.Their results suggest that the pathway of uric acid synthesis may be the same as that in birds (Buchanan, 1951), where it involves a building up of the purine ring system on a basis provided by ribose sugar phosphate, by successive reaction with glutamine, glycine, formate, glutamate, carbon dioxide, aspartate and formate, as shown schematically opposite (from Baldwin, 1963). This view receives some support from the work of Heller and Jezewska (1959) who show that uric acid synthesis in Antherea may be stimulated by the simultaneous addition of precursors like formate, ribose-5phosphate, glutamate and aspartate, in the presence of added adenosine triphosphate (ATP). ATP is required at nearly every step of purine synthesis, so that the process is an expensive one in terms of metabolic energy. But upon completion of the ring system, ribose-5’-phosphate is split off to

THE EXCRETION O F NITROGEN I N INSECTS

41

liberate hypoxanthine, and this is subsequently oxidized first to xanthine and then to uric acid. In view of the demonstration that a dehydrogenase system is involved (Irzykiewicz, 1955), it may be that a substantial proportion of energy is recaptured at this stage. The recent work of Ito and Mukaiyama (1964) provides confirmation of the role of hypoxanthine and/or xanthine as intermediaries in the metabolism of protein nitrogen. These authors show that an increase in the protein intake of silkworms causes an increase in the activity of .*

(carbon dioxide).; * a .

(formare)

8

O= -pyrophosphate

xanthine oxidase. On the basis of these results, and of the earlier findings of Hayashi (1961) that adenase activity of silkworms extracts is low compared with the guanase activity, they suggest that a pathway involving guanine and xanthine may be of importance in protein catabolism, but this does not seem to follow. It should be emphasized that evidence for the existence of a reaction sequence in insects similar to the one which has been established for birds is far from complete. Only a few of the steps have been unequivocally confirmed, and some of the results obtained (e.g. the stimulating effect of malonate and urea) cannot easily be interpreted on this basis. The possibility cannot at present be excluded that different metabolic pathways may be operative in the synthesis of uric acid by insects. C . THE FORMATION O F UREA

Urea has been reported as a minor constituent of the haemolymph (see review by Buck, 1953), of the tissues (e.g. Leifert, 1935; Brown, 1938b) and of the excreta (e.g. Powning, 1953; Razet, 1961) of many insects. Since none of the species investigated in this respect possess an active allantoicase, it seems unlikely that the urea could represent a purine breakdown product, nor can it be considered to derive from the diet in all of the species concerned. The question of its metabolic derivation in insects must therefore be considered.

42

E. B U R S E L L

One possibility is that it might be produced from amino acid nitrogen through the mediation of the ornithine cycle (see Gilmour, 1961), which has been shown to be operative in other uricotelic groups like the birds and reptiles (Cohen and Brown, 1960). Arginine, citrulline and ornithine occur as intermediaries in this reaction sequence, and all of these amino acids have been detected in certain insects (Garcia et al., 1956a); and arginase, the enzyme responsible for the hydrolysis of arginine to form ornithine and urea, has been detected in several species (Kilby and Neville, 1957; Garcia et al., 1956b; 1957 and 1958; Szarkowska and Porembska, 1959). On the other hand, arginase appears to be absent from extracts of fat body of Calliphora (Desai and Kilby, 1958a), and in Celerio euphorbiae, the ornithine cycle has been shown to be incomplete, despite the presence of its three constituent amino acids (Porembska and Mochnacka, 1964). It is natural to associate the presence of arginase with the operation of an ornithine cycle, but this may well involve a misinterpretation, as has been pointed out by Kilby (1963); it is quite possible that the function of arginase in insects is not the production of urea, but the supply of ornithine “for some as yet unknown purpose”. Direct biochemical evidence in favour of the existence of the ornithine cycle is thus very inconclusive, and the results of relevant nutritional studies are hardly less equivocal. Hinton (1955) showed that Drosophila has a developmental requirement for arginine, and that arginine can be partially replaced by citrulline, but not at all by ornithine; similar results have been obtained by Davis (1962) with larvae of Oryzaephilus. The question of the synthesis of urea in insects must clearly be left open. D. THE FORMATION OF AMMONIA

Ammonia occurs as a minor excretory product in many insects, but it is only in aquatic forms (cf. Staddon, 1955) or in meat-eating fly larvae (Brown, 1938b) that it constitutes a substantial proportion of the nitrogenous waste. The manner of its formation has not yet been satisfactorily established. Where it occurs in small quantities it is possible that it may derive entirely from nucleic acid breakdown products, by deamination of purines under the influence of adenase, guanase or adenosine deaminase, all of which enzymes have been found in insects (see above). But in species which excrete large amounts of ammonia, the nitrogen must be assumed to derive from protein sources. Attempts to demonstrate deamination of amino acids by homogenates of dipteran larvae have not been successful, nor does a diet of

THE EXCRETION OF N I T R O G E N IN INSECTS

43

free amino acids lead to an increase in ammonia production by them. A protein diet, on the other hand, causes a substantial increase in ammonia output, and homogenates are capable of causing the liberation of ammonia from various peptone mixtures (Brown and Farber, 1936; Brown, 1938b). It would seem, therefore, that the ammonia produced by feeding larvae results from a deamination of proteins or higher breakdown products, but neither the mechanism nor the precise substrate have been identified. In other insects it is likely that ammonia arises in part by direct deamination of amino acids (Bheemeswar, 1959), since amino acid oxidases have been demonstrated in the fat body of a number of insects (Kilby and Neville, 1957; Desai and Kilby, 1958a; Auclair, 1959). Alternatively, amino acids may undergo transamination with a-ketoglutaric acid as the amino acceptor, and the glutamic acid formed may be deaminated under the influence of glutamic dehydrogenase : L-amino acid a-keto acid

x

NH3

a-ketoglutaric acid

.

glutamic acid

Y (tmnsaminase)

:

NAD .H NAD

Y (gluramic dehydrogenase)

Glutamic dehydrogenase activity has been demonstrated in the cockroach Periplunetu (McAllan and Chefurka, 1961b) and in extracts from the fat body of locusts (Kilby and Neville, 1957), and the presence of transaminases has been detected in a variety of insects (Kilby and Neville, 1957; McAllan and Chefurka, 1961a, b; Zandee et al., 1958; Murphy and Micks, 1964; Chen and Bachmann-Diem, 1964; Emmerich et ul., 1965). Clearly there are a number of ways in which ammonia may be formed from protein nitrogen in insects ; but what contribution the different reactions make to the total ammonia output of a given insect cannot be decided on the basis of available evidence. E. AMINO ACIDS

Small quantities of a variety of amino acids have been found in the excreta of different insects (e.g. Harrington, 1961; Mitlin et ul., 1964), but it is possible that their presence should in many cases be interpreted as a loss of amino acid rather than as an excretion of amino nitrogen. In plant-sucking Hemiptera (e.g. Mittler, 1958) the quantities involved

44

E. B U R S E L L

are substantial in relation to the total nitrogen output, but in these insects the amino acids should perhaps be regarded as in the nature of a faecal material. In a few insects, however, large quantities of particular amino acids are excreted (Powning, 1953; Bursell, 1964a), and here there can be little doubt that a disposal of metabolic waste is involved. F. MISCELLANEOUS MATERIALS

A number of other nitrogen-containing substances have been identified as components of insect excreta; for instance, creatine has been identified in the excreta of Luciliu (Brown, 1938a) and of Rhodnius (Wigglesworth, 1931), and haematin in that of the tsetse fly (Bursell, 1964b); but in view of the diet of these insects, it seems likely that the substances represent products of digestion rather than of nitrogenous metabolism. Pteridines have been identified in the excreta of the milkweed bug, Oncopeltus fusciutus (Bartel et ul., 1958) and has been shown to make up over 5% of excretory nitrogen (Hudson et ul., 1959); but little is known of the metabolism of these substances in insects, and they will not be further discussed.

111. THEEXCRETION OF NITROGENOUS END P R O D U C T S The data presented in Section I1 have shown that enzymes capable of catalyzing the formation of a variety of end products are widely distributed among insects. But the occurrence of an enzyme which catalyses a given reaction does not necessarily mean that the product of that reaction constitutes an important component of nitrogenous excretion. Many of the substances considered may have specific parts to play in the general metabolism of the insect, and this could account for the occurrence of the corresponding enzymes. It is only by an analysis of excretory material that the contribution of different metabolic pathways to the disposal of nitrogenous waste materials can be accurately evaluated. A great deal of information is available on this point, but unfortunately it is seldom complete. The aims of investigators in this field have differed greatly. Some have been concerned simply with demonstrating the presence or absence of a specific substance in excretory material; some have focused their attention on certain classes of substance, purine degradation products for example, to the neglect of others, and so on. It is only in very few cases that the excretory material of a given insect has been accurately partitioned among all the possible nitrogenous waste products. But even with the somewhat fragmentary information which is available, it is possible to get a

THE EXCRETION OF NITROGEN I N INSECTS

45

reasonable indication of the tremendous diversity of excretory patterns which exists within the class, and the picture that emerges is not at first sight easy to reconcile with the traditional view of insects as a uricotelic group of animals. In view of the large number of species which have been investigated during recent years, it will be convenient to give a separate account of each of the orders whose members have been studied, and the classification adopted by Imms (1958) will be used for this purpose. The proportion of nitrogenous waste in the faeces of insects varies enormously, depending to a large extent on feeding habits. In herbivorous species, for instance, the nitrogen load may be light and undigested plant material often makes up the bulk of faecal matter, so that uric acid may comprise no more than 0.1% of the dry weight (e.g. Razet, 1961); in blood-sucking insects, on the other hand, the nitrogen load is heavy, there is little indigestible material, and uric acid may account for as much as 60% of the dry weight of faeces (e.g. Bursell, 1964a). In order to facilitate comparison of values which may differ by several orders of magnitude, the quantity of nitrogen excreted in different nitrogenous end products will be expressed as a proportion of nitrogen in the predominant end product. For instance, an insect which excretes 100 mg of uric acid, 10 mg of urea and 2 mg of ammonia, which would be 33.3 mg of uric acid nitrogen, 4.6 mg of urea nitrogen and 1.6 mg of ammonia nitrogen, will be listed as: Uric acid Urea Ammonia 1-00 0.14 0.05 showing that for every 100 mg of uric acid nitrogen excreted, 14 mg of urea nitrogen and 5 mg of ammonia nitrogen are disposed of. It must be emphasized that values of this kind indicate only the proportionate composition among the substances assayed. In some cases it is possible that unidentified substances, or substances not quantitatively estimated, may exceed the listed products in importance. Furthermore, values quoted for different species in the tables which follow should be regarded as no more than generally indicative of the proportionate composition in different species. Variability within species is known to be great, and in many cases determinations have been based on single samples (Razet, 1961). A . COLLEMBOLA

In view of the occurrence in a member of this order of three of the uricolytic enzymes, uricase, allantoinase and allantoicase (Razet, 1961),

46

E. BURSELL

it is a pity that no data are available on the composition of excreta, so that the extent to which degradation of uric acid is a feature of nitrogenous excretion must remain in doubt. B . ORTHOPTERA

Some of the available information concerning the nitrogenous end products in members of this order has been summarized in Table I. A number of other species have been investigated (Razet, 1961), but in TABLE I Excretory products in the Orthoptera, Odonata and Dermaptera Uric Allantoic Amino acid Allantoin acid Urea Ammonia acids

Author

Order ORTHOPTERA Schistocerca gregaria 1.00

0.00

0.00

+

+

Locusta migratoria

1-00

0.00

0.10

-

-~

MeIanopIus bivittatus

1.00

-

-

0.07

053

Acheta dornesticus

1.00

0.01

0.01

0.00

-

Mantis religiosa Periplaneta americana BIatta orientalis Carausis morosus

1-00

0.01

0.00

-

-

1.00

0.00 0.64

0.00

-

__

-

0.64 0.69

1.00

-

-

- Razet, 1961

0.44

-

-

--

Razet, 1961

Order ODONATA Aeshna cyanea (larva)

0.08

-

-

Staddon, 1959

Order DERMAPTERA ForficuIa auricuIaria 1.00

-

1-00

_.

0.00

-

1.00

-

-{

Chauvin, 1941 Razet, 1961 Razet, 1961 Nation and Patton, 1961

0.20 Brown, 1937

Razet, 1961 Nation and Patton, 1961 - Razet, 1961

-{

Razet, 1961

- Razet, 1961

the interests of brevity a selection, illustrative of the range of variation within the order, has been made for this group, as for most of the others. Uric acid is clearly the predominant end product in many Orthoptera, with other components appearing in relatively small amounts. Blattu orientalis, however, excretes most of its nitrogen as allantoic acid, while the stick insect, Curuusis morosus, has a preponderance of allantoin in its excreta. Such differences do not seem to be correlated at all

THE EXCRETION OF NITROGEN I N INSECTS

47

closely with the presence of the corresponding uricolytic enzymes. Achetu has been shown to possess active uricase, allantoinase and allantoicase,yet it excretes virtually none of the correspondingproducts; Cuurausishas a highly active allantoinase (Poisson and Razet, 1952), yet allantoin predominates in its excreta. Nor is there any apparent correlation with feeding habit, since uric acid predominates in the excreta of herbivorous, and omnivorous as well as carnivorous species. In fact, consideration of just this one order illustrates the sort of diversity in respect of nitrogenous excretion which succeeding pages will show to characterize the class as a whole. The deposition of uric acid in cells of the fat body seems to be a feature of many orthopteran species, particularly among the Blattidae (e.g. Srivastava and Gupta, 1960), and uric acid may comprise as much as 10% of the total dry weight of such insects (Razet, 1961). Haydack (1953) has shown that the amount of uric acid stored in the fatty tissues of the American cockroach is greatly affected by the level of protein intake. On a high protein diet the fat body becomes enlarged, and is filled with white deposits of uric acid. If such insects are transferred to a low protein diet the uric acid deposits largely disappear. The readiness with which uric acid may be mobilized from fat body deposits in these, as in other, insects, has led to the suggestion that it may serve in part as a reserve of nitrogen for synthetic purposes (Ludwig, 1954; Ross, 1959), and there is some indication that endocrine control mechanisms may be involved (Bodenstein, 1953). Recent work by Roth and Dateo (1964 and 1965) has shown that in males of certain cockroaches the accessory sex glands play a special role in the storage of uric acid, and that as much as 5% of the live weight of males may be attributed to uric acid deposited in the utriculi majores of these glands. Most of this uric acid is poured over the spermathecae during copulation, and the sex glands may thus be said to serve as accessory excretory organs. C . ODONATA

Excretion in the aquatic larva of Aeshna cyuneu has been investigated by Staddon (1959). In starving animals most of the excretory nitrogen appeared as ammonia, with uric acid making up less than a tenth of the ammonia nitrogen (see Table I). When the larvae were fed on protein (heat-coagulated egg white) the output of ammonia increased greatly, while uric acid excretion remained constant. It would seem that ammonia is the main end product of protein catabolism, while uric

48

E . BURSELL

acid represents the end product of purine metabolism, and is therefore unaffected by protein intake: this is in accord with the absence of uricolytic enzymes in the species investigated by Razet (1961). D . DERMAPTERA

The excreta of Forjicula has been shown to contain a high proportion of uric acid (see Table I), and despite the presence of an active allantoinase, no allantoic acid could be demonstrated (Razet, 1961). E . HEMIPTERA

1. Sub-order Heteroptera The excretory material of a number of plant bugs has been investigated, and some of the results are summarized in Table 11. Allantoic acid has not been recorded for any member of this group, and this is in TABLE I1 Excretory products in the Heteroptera Uric Allantoic Amino acid Allantoin acid Urea Ammonia acids

____

Micrelytra fossularum Verlusia rhombea Palomena prasina Rhaphigaster griseus Dysdercus fasciatiis Rhodniusprolixus

0.65

0.02 1.00 1.00

0.00 0.00

1.00 1.00

1.00 0.87

1 .oo

0.00

~

0.00 0.00 0.00

0.00 ~~

-

0.26 0.03

Author -

.__-

-

Razet, 1961 Razet, 1961 Razet, 196:

- Razet, 1961 0.24 Berridge, in press trace Wigglesworth, 1931 Harrington, 1956 and 1961

accord with the apparent absence of the corresponding enzyme, allantoinase, in the large number of species examined by Razet (1961). Allantoin, on the other hand, occurs in all species, and is the predominant end product in several, which relates well to the widespread presence of a highly active uricase among members of this suborder (Poisson and Razet, 1953; Razet, 1961). Allantoin is the predominant end product in Dysdercus, but substantial quantities of amino nitrogen have been recovered from the excreta, and urea is also an important component (Berridge, 1966);

THE EXCRETION OF NITROGEN I N INSECTS

49

in view of the general absence of allantoinase and allantoicase which characterizes the group as a whole, the urea is unlikely to represent a product of uricolysis, and the possibility of faecal contamination can be excluded in this insect, since the alimentary canal is discontinuous. Urea has also been found in the excreta of Oncopeltus fasciatus, but in this species uric acid constitutes the predominant end product (Nation and Patton, 1961). The blood-sucking hemipteran RhochiusproZixus is a member of the suborder, and its excretory metabolism has been the subject of several studies (Wigglesworth, 1931; Harrington, 1956, 1961). Uric acid is the main end product, but urea accounts for 2.3% of the dry weight of excreta, and a number of amino acids have been detected together making up 0.2% of the total dry weight, with histamine and histidine predominant among them. In view of the fact that the food of this insect is composed almost entirely of amino acids, arising by hydrolysis of blood proteins, the appearance of such very small quantities of amino acid nitrogen in the excreta should perhaps be interpreted as a loss, rather than as an excretion, of amino acids. 2. Sub-order Homoptera Investigations of excretory metabolism in this group of insects has centred upon the occurrence of large amounts of amino acid in the honeydew of various aphids and coccids (see review by Auclair, 1963). These substances account for 13.2% of the dry weight of honeydew in Myzus circumflexus (Maltais and Auclair, 1952) and make up the bulk of nitrogenous material also in Brevicoryne brassicae (Lamb, 1959). The pattern of amino acids in honeydew is qualitatively similar to that of the plant juices ingested, though the concentration may be slightly lower due to absorption of nitrogenous material (Mittler, 1958 with Tuberolachnus salignus), or somewhat higher (Auclair, 1958 with Acyrthosiphon pisum) possibly due to a concentration of fluids in the insect associated with evaporative losses of water. But it would seem that the insects are ingesting nitrogenous substances in excess of their requirements, and that this excess is voided as a faecal material. If this is the correct interpretation then such amino acids have not been subjected to metabolic manipulation, and should hardly be considered as end products of nitrogenous metabolism. Gray and Fraenkel(l954) have reported the occurrence of uric acid in Pseudococcus citri on the basis of qualitative, colour reactions, but Mittler (1958) failed to find evidence of uric acid or of ammonia in Tuberolachnur.In Breuicornis neither uric acid nor urea, allantoin or

50

E. BURSBLL

allantoic acid could be detected (Lamb, 1959), but ammonia was found to constitute 0.5% of the total nitrogen in honeydew. In view of the rapid throughput of liquid in these insects it is not impossible that even such low concentrations of ammonia may represent a high proportion of truly excretory nitrogen. Uricase has been demonstrated in homogenates of Aphis brassicae, but other members of the group which have been investigated appear to be devoid of uricolytic enzymes (Razet, 1961). F. C O L E O P T E R A

Table I11 summarizes results of investigations of the excretory metabolism in beetles. Uric acid is the predominant end product in three of the species examined, and in these uricolytic activity has been shown to be absent or feebly developed (Razet, 1961). In Chrysobothris TABLE I11 Excretory products in the Coleoptera and Neuroptera Uric Allantoic Amino acid Allantoin acid Urea Ammonia acids Author Order COLEOPTERA 1.00

0*00

0.00

-

-

-

1.00 1.00

0.01 0.10

0.03 0.00

-

-

- Razet, 1961 Razet, 1961

0.83

1.00 -

0.00 -

-

0.72

1.00

0.57

- Razet, 1961 0.50 Pomhg, 1953

Order NEUROPTERA Uroleon nostras (larva) 1.00

0.03

0.00

-

-

Melolontha vulgaris Chaetocarabus intricatus Procrustes coriaceus Chrysobothris afinis Attagenuspiceus

Razet, 1961

-

- Razet, 1961

the quantity of allantoin recovered in excreta exceeds that of uric acid, which suggests the presence of an active uricase, but this has not been directly verified. In the carpet beetle, Attagenus, urea is the predominant end product identified, substantial quantities of ammonia are excreted, and the presence of the sulphur-containing amino acid, cystine, accounts for a considerable proportion of excretory nitrogen (Pawning, 1953); its occurrence is correlated with the high cystine content of keratin, which forms the major component of diet in this species, and it is possible that this amino acid should be considered to represent a faecal material rather than an excretory product. It must be noted

THE EXCRETION O F NITROGEN I N INSECTS

51

that the nitrogenous components listed for this insect make up only a proportion of the total nitrogen excreted, and some unidentified excretory products must be involved. The excreta of the mealworm beetle, Tenebrio molitor, contains mostly uric acid, but the presence of urea and allantoin has been established (Nation and Patton, 1961) although none of the uricolytic enzymes could be demonstrated in this species (Razet, 1961). Urea and ammonia have been found as constituents of the excreta of other members of the order (Delaunay, 1931) and ammonia has been identified in the excreta of Ephestia kuhniella (Payne, 1936). The excreta of the boll weevil, Anthonomus grandis has been examined for the presence of non-protein amino acids, and these were found to constitute 3.2% of the total nitrogen, with about a third attributable to free amino acids (Mitlin et al., 1964). Storage of uric acid in cells of the fat body has been reported for one of the species of beetle examined by Gupta and Sinha (1960), but storage excretion does not appear to be a general characteristic of the group, to judge by the generally low uric acid contents recorded by Razet (1961). G . NEUROPTERA

Data for the larva of Uroleon have been included in Table 111. Since the alimentary canal is discontinuous, excretory material could be collected free of faecal contamination. 33% of the dry weight was uric acid, and only a small quantity of allantoin was present; allantoic acid could not be detected. In the larva of Chrysopa carnea the deposition of uric acid in cells of the fat body has been reported (Spiegler, 1962). The presence of large quantities of ammonium bicarbonate in excretory fluids of the aquatic larva of Sialis lutaria was demonstrated by Shaw (1955). The excretory metabolism of this insect was further investigated by Staddon (1955) who showed that 86% of the nitrogen output of starving animals appeared in the form of ammonia. H . HYMENOPTERA

The excreta of certain herbivorous larvae belonging to this group has been investigated by Razet (1961), and some of his results are summarized in Table IV. Here again some species excrete predominantly uric acid, in others allantoin or allantoic acid may be important constituents, and in some they achieve dominance. The differences between

52 E. BURSELL species cannot be correlated with differencesin host plant, nor do they accord with the distribution of uricolytic enzymes. Uricase is poorly developed in both the genera listed; and Pteronidea sulicis, which has a highly active allantoinase, excretes a very small proportion of allantoic acid, while in P. ribesi allantoinase activity is slight, but allantoic acid forms a high proportion of the excretory material. TABLE IV Excretory products in the Hymenoptera and Diptera Uric Allantoic Amino acid Allantoin acid Urea Ammonia acids Author Order HYMENOPTERA Hemichroa alni

1.00

0.00

0.00

- Razet, 1961

1.00

0.03

0.06

- RBWt, 1961

0.78

1.00

0.90

- R m t , 1961

1.00 1-00 1.00

0.04

0.30 0.30

0.00 0.62

0.05

1.00

(larva) Pteronidea salicis

(larva) Pteronidea ribesi

(larva) Order DIPTERA Compsilura concinnata Tipulapaludosa Lucilia sericata Lucilia sericata Oarva) Lucilia sericata

(pupa) Bibio marci (larva) Aedes aegypti Anopheles guadrimaculatus Culex pipiens Glossina morsitans

I

0.45

1*0° 1.00

-

-

0-30

0.02

-

1.00

0.00 1.00

-

0.15

0-36

-

0.22

+

0.18

- Razet, 1961 - X m t , 1961

-

-{

Brown, 1936; 1938a and b

-

- Razet, 1961

0.11 Irrevere and Terzian, 1959 0.22 Bursell, 1964b

I . DIPTERA

A number of insects belonging to this order have been studied, and results are included in Table IV. Most of the adults examined are predominantly uricotelic, though in Tipula, where active uricolytic enzymes have been demonstrated (Razet, 1961), substantial amounts of allantoin and allantoic acid are present in the excreta. In the sheep ked, Melophugus ovinus, uric acid is the main end product, but quantities of xanthine and hypoxanthine may appear in the excreta of pregnant females (Nelson, 1958).

53 The excretory metabolism of several blood-sucking members has been the subject of detailed investigation. Uric acid is the dominant excretory product in mosquitoes, but urea and ammonia occur in fairly high concentration, and amino acids make up a substantial proportion of total nitrogen; there is little difference between the three species examined, and in Table IV results have been averaged. The effect of nutrition on the excretion of Aedes aegypti has been studied by Terzian and his colleagues (1957), who show that on a diet of sugar there is a decrease in the nitrogen output, and the proportion of uric acid in the excreta falls to about 4%. Following a blood meal the nitrogen output increases greatly, and the proportion of uric acid rises to SO%, thus confirming that uric acid is the end product of protein metabolism. In the tsetse fly also, the predominant excretory product is uric acid; urea occurs in trace amounts, and a substantial proportion of nitrogen is excreted in the form of histidine and arginine. These two amino acids, which together make up about 10% of the protein amino acids in human blood, are particularly rich in nitrogen (see Fig. 1). It would presumably be uneconomical to deaminate materials of this kind, since any benefit which the insect might derive from deamination products would be offset by metabolic losses involved in uricotelic detoxication of the nitrogen they contain. In view of this, a quantitative elimination of ingested arginine and histidine might be expected, and this is roughly in accord with observation (Bursell, 1964b). There is some evidence that all of the amino acids liberated during digestion are absorbed from the midgut, and that arginine and histidine are subsequently excreted, so that they cannot be regarded simply as faecal materials. Indeed, recent work has shown that arginine becomes rapidly labelled following injections of C14-glutamate(E. Bursell, unpublished), and it is possible that this substance may play a more active role in excretion than was originally envisaged. The occurrence of small quantities of ammonia has been reported in the excreta of adult Diptera by a number of workers (Brown, 1938b; Lennox, 1940; Hitchcock and Haub, 1941; Sedee, 1958), but it is only in the meat-eating larvae of species like Calliphora and Lucilia that this product becomes dominant (Brown, 1936; 1938a, b; Robinson, 1935; Robinson and Baker, 1939). Allantoin and uric acid are present in relatively small amounts in the larval excreta of Luciliu (see Table IV), and the proportion between them appears to depend in part on larval diet; with larvae fed on casein allantoin predominates, but on a meat diet relatively little allantoin is produced (Brown, 1938a). THE EXCRETION O F NITROGEN I N INSECTS

54

E. B U R S E L L

Allantoin is an important constituent of excreta in larvae of Bibio, and allantoic acid occurs as well, in accord with the presence of an active allantoinase in larval extracts (Razet, 1961). The excretory products which accumulate in the meconium during pupal development have been examined by Brown (1938a, b) in Lucilia. A considerable amount of ammonia was found, but uric acid constitutes the bulk of excretory nitrogen and no allantoin could be detected. The absence of allantoin is correlated with a loss of uricase activity at this stage of the life history; the enzyme disappears suddenly when larvae leave the meat, to reappear immediately after emergence from the pupa. The results of investigations of dipteran excretion provide a striking illustration of the lability which appears to characterize this aspect of metabolism, with the distribution of nitrogenous end products varying widely from stage to stage in the life history of a single species. J. LEPIDOPTERA

A great deal of information is available concerning the uricolytic metabolism of this group, thanks to the work of Razet (1961). A selection of results has been summarized in Table V, where the proportions of uric acid, allantoin and allantoic acid in the excreta of different stages in the life history of members of the order have been set out. Variability within the group is considerable; many of the adults are predominantly uricotelic, but allantoin and allantoic acid occur as minor components in several species, and in some allantoic acid is the predominant end product. In pupal stages uric acid is predominant in all the species listed, including those whose adults show a preponderance of allantoic acid; but the proportions of allantoic acid and allantoin are often appreciable, and in some species exceed those found in adult excreta (e.g. Aglais urticae). In larval stages the three earcretory products are more evenly distributed in the excreta of different species; where allantoic acid is the end product, uric acid occurs in substantial proportion, and vice versa. But there is little correlation between adult and larval excretory metabolism, for some of the species whose adults are predominantly uricotelic show a preponderance of allantoin in larval excreta, while some of the species whose adults have allantoic acid as the main end product have uricotelic larvae. In certain species it has been demonstrated that larval diet may have an important effect on excretory metabolism. For instance, with

55

THE EXCRETION O F NITROGEN I N INSECTS

larvae of Lasiocampa trifolii fed on leaves of Hordeum murinum the predominant excretory product is allantoic acid, while a diet of Betulus aZba produces a predominance of uric acid. Such effects cannot be attributed to the pre-existence of nitrogenous end products in the diet, since of all the host species examined only one, the cabbage Brassica TABLE V Excretory products in the Lepidoptera (Razet, 1961) Uric acid

1. ADULTS Mammestra brassicae Agritos comes Aglais urticae Trigonophora meticulosa Pieris brassicae Vanessa atalanta Conistra vaccinii 2. PUPAE M . brassicae A. comes A . urticae T. meticulosa P. brassicae V. atalanta C. vaccinii

3. LARVAE M . brassicae A. comes A. urticae T. meticulosa P. brassicae V. atalanta C. vaccinii

1.00 1.oo 1.oo 1.00 1.00 0.89 0.12

1.00 1.00 1.00 1.00

Allantoin -

0.00 0.01 0.04 0.14 -

-

0.29 -

1.oo 1.00 1.00

0-03 0.14 0.04

0.49 0.67 0.56 1-00 0.28 1.oo 1.00

0.11 0.02 0.18 0.02

0.16 0.26 0.06

Allantoic acid 0.00 0.01

0.06 0.05 0.01

1.00 1.00

0.03 0.01 0.43 0.19 0.05 0.29 0.04 1.00 1.00 1.00 0.12 1.00 0.20 0.05

oleracea, could be shown to contain small quantities of allantoin and allantoic acid. The occurrence of uricolytic products in the excreta of different members of the order is in general accord with the presence, in most species, of the corresponding uricolytic enzymes, but in point of detail there is little agreement. In some species where enzymatic activity is 34-A.I.P.

4

56

E. BURSELL

relatively high the corresponding end product is poorly represented in the excreta (e.g. allantoinase in adults of Aglais urticae), while species with relatively low enzymatic activity may show a preponderance of the reaction product in their excreta (e.g. Conistra vaccinii). The excretory metabolism of lepidopteran larvae at different stages of larval life has also been investigated by Razet (1961), and the results serve further to emphasize the lability of this function within the life history of the individual, for the ratio of allantoin to uric acid may fluctuate enormously from day to day. In most of the species studied there is a general tendency for uric acid to become increasingly important with larval age. A number of end products other than these three have been assayed in the excreta of different members of the order. In Bombyx mori, amino acids have been demonstrated in excretory material of the larva (Yoshitaka and Aruga, 1950); uric acid is the predominant end product in this species, and the quantity exsreted has been shown to increase on a diet rich in protein (It0 and Mukaiyama, 1964). In larvae, of Corcyra cephalonica uric acid is again predominant in the excreta, but small quantities of urea and of xanthine have been demonstrated (Srivastava, 1962) ; both xanthine and hypoxanthine have been identified in the excreta of the wax moth, Galleria melonella (Nation, 1963; Nation and Patton, 1961), with hypoxanthine making up as much as 10% of the total; ammonia has also been identified in the larval excreta of this species (Zielenska, 1952). In the clothes moth, TineoZa bisselliella, uric acid also predominates, but for every 100 mg of uric acid nitrogen excreted, 24 mg of ammonia nitrogen and 10 mg of urea nitrogen are disposed of. In addition, the excreta of this species contains a substantial amount of amino nitrogen in the form of cystine, which correlates with its diet of keratin (Waterhouse, 1952; Powning, 1953). I V . CONCLUSIONS The preceding pages have given some indication of the tremendous progress which has been made in the field under review during the last 15 years. A number of uricolytic enzymes have been investigated in tissue homogenates, their distribution among Werent tissues has been established and some of their properties have been described. A great deal has been discovered about xanthine oxidase, or xanthine dehydrogenase as it should more properly be called, and here studies have advanced to the stage of enzyme purification. But unfortunately there remain several gaps in our knowledge of nitrogenous metabolism

THE EXCRETION OF N I T R O G E N I N INSECTS

57

at the level of enzyme studies. The contribution made by different reactions (adenase and guanase deamination, oxidative deamination of amino acids, the glutamic dehydrogenase system) to the appearance of ammonia remains under dispute. The origin of urea is still uncertain, and, above all, the precise pathway of uric acid synthesis has not been unequivocally settled. The evidence concerning these aspects of nitrogenous metabolism is often contradictory, and the problems stand in urgent need of resolution. The application of radioactive tracer techniques, coupled wit% the use of insects whose nutrition, in terms of protein and nucleoprotein intake, can be rigidly controlled, would seem to offer prospects of rapid advance in this field. The role of urea in nitrogenous excretion has never been fully understood; it is present in the excreta of many insects, but usually forms a rather minor component. It cannot be regarded as a purine degradation product, since so far no insect has been shown to possess the enzymes required for the degradation of uric acid to this stage. The presence of arginase in many insects suggests that it may originate by hydrolysis of arginine, but evidence for the existence of the complete omithine cycle is weak. It seems possible that arginine may be the raw material for its production, but that its occurrence in insects should be seen as a reflection of the use of arginine as a phosphagen in this class of animals (Razet, 1964) ;the precise implications of this suggestion have, however, yet to be worked out. The data summarized in Section I1 of this review give some indication of the complexities involved in the excretory metabolism of insects. In the face of such complexities one may be forgiven for looking back with a degree of nostalgia to the early generalization-that insects as a class are uricotelic, and that the production of uric acid as the main end product of nitrogenous metabolism can be seen as a reflection of the shortage of water to which most members of the group are subject. This shortage of water would make toxic substances, like ammonia, and%oluble substances, like urea, unsuitable as waste products. It was possible to accommodate certain exceptions within the general framework of this generalization; for instance, the predominance of ammonia as an excretory product of aquatic larvae, of certain dipteran larvae and perhaps of certain plant-sucking bugs, could be interpreted on the basis of the ready availability of water associated with these modes of life. Similarly, the excretion of substantial proportions of specific amino acids could be seen as an adaptation to special diets; insects which ingest a lot of keratin excrete a lot of the sulphur-containing amino acid which this diet contains; blood-sucking insects, whose

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diet contains a high proportion of nitrogen-rich amino acids, may eliminate these unchanged, though why this should occur in the tsetse fly and to some extent in mosquitoes, but not in Rhodnius, is unclear. Up to this point it would appear permissible to retain the view of uricotelism as an insect characteristic. But what investigations of the last ten years have brought to light is the high proportion of purine derivatives other than uric acid that occur in the excreta of many insects. These include not only degradation products like allantoin and allantoic acid, but also some of the reduced derivatives like xanthine and hypoxanthine. Since these substances may occur in the excreta of insects in amounts far exceeding any that could be attributed to nucleic acid catabolism, they must derive largely from amino acid nitrogen. The situation, therefore, seems to be that amino acid nitrogen is detoxicated by incorporation in a purine ring system at the expense of considerable amounts of metabolic energy; and that the purine ring system is then broken down in a sequence of hydrolytic reactions which are unproductive of metabolic energy, and lead to the formation of reaction products that differ appreciably from uric acid neither in respect of solubility nor in respect of nitrogen content. Coupled with the apparent lack of correlation between the proportion of degradation products in the excreta of different insects and their mode of nutrition and way of life, this provides small basis for an interpretation in terms of water balance or of excretory efficiency. In seeking for an alternative basis of interpretation, perhaps the most sigmficant finding is the localization of enzyme activity in cells of the Malpighian tubules and of the midgut, both of which are regions which have been implicated in the elimination of excretory material. It seems possible that the degradation of the purine ring may in some way be associated with the transfer of material across a secretory epithelium (M. J. Berridge, in the press), perhaps by some form of facilitated diffusion. The ability to break the purine ring to the stage of allantoic acid appears to be widely, and maybe universally, distributed within the class. It is true that negative results of enzyme assays have been reported for a number of insects (Razet, 1961), but this may mean not that the enzymes are absent, but that they are present in concentrations too low to be detected by the methods used. The nature of the results obtained by Razet does not allow a great deal of reliance to be placed on negative results. In many cases the quantity of substrate broken down is inversely related to the amount of enzyme present, and Razet suggests the possibility that inhibitory factors may be involved, coming into play at high homogenate concentrations. Similar effects could be involved in

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extracts from species showing no detectable activity; indeed, the development of more refined methods led to the detection of uricase in species which had previously been thought devoid of activity (Razet, 1964). It would seem that investigations with dialysed enzyme preparations would be required for unequivocal demonstration of the absence of uricolytic activity. These considerations suggest that the development of uricotelism in insects has not been associated with a loss of uricolytic enzymes. The ability to synthesize these enzymes appears to have been preserved in the genotype, perhaps because certain of the reaction products have special parts to play in general metabolism. It is the degree to which this ability is reflected in the phenotype which is subject to immense variation from order to order within the class, from species to species within the orders, from stage to stage in the life history of a species, and from time to time within a given stage. The main problem which confronts the investigator today is to discover what is the biological significance of these variations, and with what aspect of species biology they are correlated. The demonstration that different nitrogenous end products may achieve dominance in the excretory metabolism of different insect species calls, however, for some reassessment of the generalization that terrestrial insects as a group are uricotelic. Either this statement must be qualified to take account of recent discoveries, or it must be abandoned in favour of a more complex statement of the situation. The second alternative is the one favoured by P. Razet (in the press) in his recent review of nitrogenous metabolism in insects. He proposes that insects should be assigned to one or other of a number of different categories depending on the nature of the predominant excretory end product. The terms uricotelic, ureotelic and ammonotelic would be retained to indicate a preponderance in the excreta of uric acid, urea and ammonia respectively, and two new categories would be added : allantoinotelic for insects which excrete mainly allantoin, and allantoicotelic for insects which excrete mainly allantoic acid. Apart from the rather cumbersome nature of these new epithets, a classification of this sort does not seem very helpful at the present stage. In the first place it bears a largely random relation to the generally accepted taxonomy of the class, and representatives of a given category may occur equally among primitive and advanced members. Secondly, the proposal to classify species on the basis of the predominance of a given end product would run into difficulty in the case of certain Diptera in which the larva is ammonotelic and the adult uricotelic; or

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certain Lepidoptera, which may be uricotelic in the larval stage and allantoicotelic as adults, or whose larvae may switch from one category to another depending gn age. What is described would here cease to be a species characteristic; the terms would become descriptive simply of a particular moment in the life history, and would thus be deprived of their generally accepted connotations. Even if it should be found desirable to institute a new system of classification of the kind proposed by Razet, the time to apply it to the orders of insect would not be now. For while the information available is reasonably extensive in respect of certain groups (e.g. the Lepidoptera), it is very meagre in respect of others (e.g. Collembola; Dermaptera), and these latter might have to be assigned to their place in classification on the basis of a single species. In view of the wide spectrum of end product predominance discovered in all the groups which have been thoroughly studied, it seems certain that any scheme proposed would be shortlived and subject to continual adjustment in the light of new information. For these reasons it seems to the present reviewer that some attempt should be made to retain the older generalization for the time being, if this can be done by suitable qualification. Granted the complexities reviewed in Section 11, it still remains true to say that uric acid is an important excretory product in nearly all terrestrial insecfs, and to this limited extent the statement that uricotelism is a characteristic of the class remains valid. It is not always the predominant excretory product, but where it is not, its place seems invariably to be taken by allantoin or allantoic acid, as far as present information goes. Since these substances differ from uric acid in neither of the properties which are chiefly relevant to the disposal of nitrogenous waste, namely nitrogen content and solubility, it would seem permissible to regard them all as belonging, to a single class of excretory substance, and to widen the definition of uricotelism accordingly-to regard an animal as uricotelic if it excretes uric acid or one of its primary degradation products, allantoin or allantoic acid, or some mixture of these three substances, as the predominant waste product. With such a qualification the view of insects as a uricotelic class, in line with thsir terrestrial mode of life, remains tenable, and the centre of interest in the field of nitrogenous metabolism then shifts from further cataloguing of the relative proportions of these end products in the excreta of insects to what seems at present a more rewarding problem, namely a study of the biological significance of such differences in the proportion of end products, possibly in relation to the mechanism of excretion. The most promising approach to this study, and through it to the general problem of the distribution of

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nitrogenous end products, would seem to lie with the species which show a pronounced shift in excretory metabolism during the course of their life history. Here there would be some possibility of relating observed changes in the proportion of different end products to the level and type of nutrition and to the availability of water, and in this way to gain some insight into the significance of such changes. The work of the last decade has adequately demonstrated the diversity and complexity of excretory metabolism in the insects as a whole. What seems to be required now are intensive studies of selected species in which the problem of excretory metabolism can be set squarely in the context of species biology.

ACKNOWLEDGEMENTS My sincere thanks are due to Dr. P. Razet for his kindness in presenting me with a copy of his thesis, and for allowing me to read and to refer to his review of nitrogenous metabolism before its publication.

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Prosser, C. L. (ed.) (1952). ‘‘Comparative animal physiology”. Saunders, Philadelphia. Rota, C. D. (1961). Enzymes in the hemolymph of the mealworm Tenebrio molitor L. J. N . Y. ent. SOC.69, 59-67. Razet, P. (1952). Catabolisme des purines chez les Collembole Xenyllu welchii Folsom (Insecte, Aptdrygote). C . r. Sdanc. SOC.Biol. 234, 25662568. Razet, P. (1953). Recherches sur la localisation des enzymes uricolytiques chez les insectes. C. r. Sdanc. SOC.Biol. 236, 1304-1306. Razet, P. (1954). Sur l’dlimination d’acide allantoique par quelques insectes Ldpidopteres. C. r. Sdanc. SOC.Biol. 239, 905-907. Razet, P. (1956). Sur l’dlimination simultanke d‘acide urique et d‘acide allantoique chez les insectes. C . r. Sianc. SOC.Biol. 243, 185-187. Razet, P. (1957). L’uricolyse chez les insectes. Arch. Orig. Serv. Docum. C.N.R.S., 361. Razet, P. (1961). Recherches sur l’uricolyse chez les insectes. These Doct. Sc. Nat., Imprimerie Bretonne, Rennes. Razet, P. (1964). Le probleme de l’acide urique chez les arthropodes antennates. 12th Znt. Congr. Ent. 220-221. Razet, P. (in the press). Les elements terminaux du catabolisme azote chez les insectes. Annke biol. Robinson, W. (1935). Allantoin, a constituent of maggot excretions, stimulates healing of chronic discharging wounds. J. Parasit., 21, 354-358. Robinson, W. and Baker, F. C. (1939). The enzyme urease and the occurrence of ammonia in maggot-infected wounds. J. Parasit. 25, 149-155. Robinson, W. and Wilson, G. S. (1939). Changes in the concentration of urease during pupal development of the blowfly Phormia regina. J. Parasit. 25, 455459. Rocco, M. L. (1936). Prdsence de l’allantoinase chez les insectes. C. r. Sdanc. SOC.Biol. 202, 1947-1948. ROCCO,M. L. (1938). Le mdtabolisme des composds d’origine purique chez les insectes. C . r. Sdanc. SOC.Biol. 207, 1006-1008. Roeder, K. D. (ed.) (1953). “Insect Physiology”. John Wiley and Sons, New York. Ross, D. J. (1959). Changes in the activity of uricase and xanthine oxidase during the life cycle of the Japanese beetle, Popillia japonica Newm. Physiol. Z00“l. 32, 239-245. Roth, L. M. and Dateo, G. P. (1964). Uric acid in the reproductive system of males of the cockroach Blatalla germanica. Science, N . Y. 146, 782-784. Roth, L. M. and Dateo, G. P. (1965). Uric acid storage and excretion by accessory sex glands of male cockroaches. J. Insect Physiol. 11, 1023-1030. Sedee, J. W. (1958). Dietetic requirements and intermediary protein metabolism of an insect (Calliphora erythrocephala Meig.). Entomologia exp. appl. 1, 38-40. Shaw, J. (1955). Ionic regulation and water balance in the aquatic larva of Sialis lutaria. J. exp. Biol. 32, 353-382. Smith, K. D., Ursprung, H. and Wright, T. R. F. (1963). Xanthine dehydrogenase in Drosophila: Detection of isozymes. Science, N. Y. 142, 226-227. Spiegler, P. E. (1962). Uric acid and urate storage in the larva of Chrysopu carnea Stephens (Neuroptera, Chrysopidae). J. Insect Physiol. 8, 127-1 32.

THE EXCRETION O F NITROGEN I N INSECTS

67

Srivastava, P. N. (1962). Physiology of excretion in the larva of Corcyra cephaIonica Stainton (Lepidoptera, Pyralidae). J. Insect Physiol. 8, 223-232. Srivastava, P. N. and Gupta, P. D. (1960). Excretion of uric acid in Periplaneta americana L. J. Insect Physiol. 6, 163-167. Staddon, B. W. (1955). The excretion and storage of ammonia by aquatic larvae of Sialis lutaria (Neuroptera). J. exp. Biol. 32, 84-94. Staddon, B. W. (1959). Nitrogen excretion in nymphs of Aeshna cyanea (Mull.) (Odonata, Anisoptera). J. exp. Biol. 36, 566-574. Stobbart, R. H. and Shaw, J. (1964). Salt and water balance: excretion. In “Physiology of Insecta”, (M. Rockstein, ed.) Academic Press, New York and London. Szarkowska, L. and Porembska, Z. (1959). Arginase in Celerio euphorbiae. Acta biochim. pol. 6,273-276. Terzian, L. A., Irreverre, F. and Stahler, N. (1957). A study of nitrogen patterns in the excreta and body tissues of adult Aedes aegypti. J. Insect Physiol. 1, 221-228. Tomita, M. and Kumon, T. (1936). Zur Chemie der Fliegen-larven. HoppeSeyler’s Z. physiol. Chem. 238, 101--104. Truszkowski, R. and Chajkinowna, S. (1935). Nitrogen metabolism of certain invertebrates. Biochem. J. 29, 2361-2365. Ursprung, H. and Hadorn, R. (1961). Xanthinedehydrogenase in Organen von Drosophila melanogaster. Experientia, 17, 230-23 1. Wagner, R. P. and Mitchell, H. K. (1948). Enzymic assay for studying the nutrition of Drosophila melanogaster. Archs Biochem. 17, 87-96. Waterhouse, D. F. (1952). Studies on the digestion of wool by insects. IV. Absorption and elimination of metals by lepidopterous larvae, with special reference to the clothes moth, Tineola bisselliella (Humm.). Aust. J. scient. Res. (B), 5, 143-168. Wigglesworth, V. B. (1931). The physiology of excretion in a blood-sucking insect, Rhodnius prolixus (Hemiptera, Reduviidae). J . exp. Biol. 8, 41 1-451. Wigglesworth, V. B. (1950). “Principles of Insect Physiology.” Methuen, London. Yoshitaka, N. and Aruga, H. (1950). Studies on the amino acids in the silkworm. IV. On the free amino acids in the silkworm faeces. J. seric. Sci., Tokyo 19, 536. Zandee, D. L., Nijkamp, H. J., Roosheroe, L., De Waart, J., Sedee, P. D. J. W. and Vonk, H. J. (1958). Transaminations in invertebrates. Archs int. Physiol. Biochim. 66, 220-227. Zielenska, Z. M. (1952). Studies on the biochemistry of Galleria melonella. Nitrogen metabolism of the larva. Acta Biol. exp. Vars. 16, 171-186.

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Lipid Metabolism and Function in Insects LAWRENCE I. GILBERT Department of Biological Sciences, Northwestern University, Evanston, Illinois, U.S.A. I. Introduction . A. General . B. Definition and classification 11. Lipid Content . A. Expression of data . B. Alterations during metamorphosis . C. Nature of insect lipids . III. Lipid Utilization . A. Digestion and absorption . B. Lipid release and transport C. Extra-digestive lipases . D. Fatty acid catabolism . IV. Lipid Biosynthesis . A. General mechanism of fatty acid synthesis B. Fatty acid biosynthesis in insects C. Phospholipid and triglyceride . D. Fatty acids in nutrition . E. Substrate interconversion . V. Hydrocarbons and Waxes A. Cuticle B. Extra-cuticular . VI. Isoprenoid Compounds . A. Nutritional studies . B. Isoprenoid biosynthesis . C. Isoprenoid content . D. Sterol modification . E. Function . F. Insect hormones VII. Conclusion . References .

.

.

. . . . . . .

.

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

70 70 71 71 71 81 89 97 97 102 110 116 127 127 130 134 145 147 152 152 155 157 157 161 168 170 175 176 186 187

In the course of this paper, the reader will encounter the following abbreviations: ADP-adenosine diphosphate; ATP-adenosine triphosphate; CoA or COASHcoenzyme A; Co@-coenzyme Q ; CTP4ytidine triphosphate; DGLAiglyceride; FAD-flavine adenine dinucleotide; FFA-free or unestersed fatty acids ; MGLmonoglyceride; NAD-nicotinamide adenine dinucleotide; NADP-nicotinamide PTC-phosphatidylcholine; adenine dinucleotide phosphate; PL-phospholipid; FTE-phosphatidylethanolamine ;TGL-triglyceride. 69

70

LAWRENCE I . GILBERT

I. INTRODUCTION A . GENERAL

The awarding of the Nobel prize in medicine to Bloch and Lynen for their contributions to the study of lipid metabolism is a heartening occurrence for many biologists and biochemists. This act pointed out once again the importance of the study of lipids, their synthesis and catabolism. It was in fact in this research area that the first definitive data were obtained on the dynamics of any protoplasmic constituents when Schoenheimer and Rittenberg (1936) studied fatty acid turnover utilizing heavy water. One prime impetus to the study of lipid biochemistry in insects in recent years has been the finding that many, if not all of the insect growth hormones, pheromones and sex attractants are lipoidal (cf. Gilbert, 1964). An understanding of the biosynthetic pathway and means of catabolizing these humoral and air-borne messengers necessitates a vigorous experimental approach to the fields of insect lipid chemistry and biochemistry. As will be discussed subsequently, lipids are also of vital importance to many insects as substrates for embryogenesis, metamorphosis and flight. And several members of this large category of biochemical compounds are without peers as vital nutritional growth factors. The purpose of this review is not so much to tabulate all knowledge of insect lipid biochemistry as to point out the gaps in our knowledge, in an attempt to attract others to this increasingly important research area. When this paper was begun, the last tome on insect lipids was more than a decade out of date (Scoggin and Tauber, 1950). In recent years however, several reviews on aspects of insect lipid chemistry and biochemistry have appeared (Clayton, 1964; Fast, 1964; Gilby, 1965 and Tietz-Devir, 1963),while other reviews that consider the metabolism of lipids along with other biochemicals have also been published (Chefurka, 1965; Gilbert and Schneiderman, 1961a; Gilmour, 1961; Kilby, 1963). Numerous papers on the biochemistry of lipids in microorganisms, fowl and mammals have appeared in the recent past and will be alluded to when considered important to the present discussion. It is of interest to note that several journals dealing exclusively with lipid biochemistry have appeared in the last eight years (Journal of Lipid Research, Lipid section of Biochimica Biophysica Acta, Steroids, as well as Advances in Lipid Research) but one has difficulty finding more than a dozen publications considering insect material in the cumulative

L I P I D METABOLISM A N D F U N C T I O N I N I N S E C T S

71

editions of these journals. It may be that as many reviews have appeared on this subject in recent years as breakthroughs in this research area. We shall concentrate on developments within the past ten years but will discuss older literature when applicable to the historical development of the topic or when it buttresses the argument. However, not all of the older literature will be discussed. Some aspects of the topic will be treated in a cursory manner, some omitted and others dwelled upon at length. The hierarchy of discussion is in part due to the author’s own interests and it is possible that topics not treated in depth (e.g. nutrition) are of more importance than those treated extensively. A final point worthy of note is that few generalizations can be made regarding all members of this largest animal class. In most cases, sophisticated experiments have been conducted on only a few species of “domesticated” insects and one cannot extrapolate the results to all insects. The multitude of different ecological niches and behavioural characteristics are no doubt reflected in a great number of metabolic variations on perhaps more than one basic theme. Notwithstanding the above, the past five years have brought us a clearer understanding of at least some aspects of the lipid biochemistry of insects. Before considering these advances it is of importance to agree on a vocabulary. B. DEFINITION A N D CLASSIFICATION

There has as yet been no rigorous definition of the term “lipid” that has been accepted categorically by all workers in the field. Generally, all compounds that are insoluble or only sparingly soluble in water and are extractable with, or soluble in, organic solvents such as diethyl ether, chloroform, acetone, etc., are termed lipids. For our present purposes, “lipids” will be defined as all compounds falling into the classification outlined in Table I, which is adapted and modified from similar tables by numerous authors, but notably Deuel (1951) and Strickland (1963). 11. LIPIDCONTENT A . EXPRESSION OF D A T A

One of the fundamental questions concerning the role of lipids in the physiology of the insect concerns the quantity of lipid contained upon, and interior to, the rigid exoskeleton. The exact amount will of course vary with physiological state, but also appears to vary according to the method of extraction and whether the resqlts are expressed in terms of dry weight, wet weight or lean dry weight. At first glance it

72

LAWRENCE I . GILBERT

would appear most desirable to express the quantity of lipid in terms of dry weight of the insect or tissue since this value can be obtained easily by drying the tissue to constant weight at about 100” and subsequently extracting the lipid. Unfortunately this procedure usually TABLE I Classification of lipids

A. Simple Lipids 1. Glycerol esters of fatty acids (mainly long-chain fatty acids; commonly called “neutral fat ”; includes monoglycerides, diglycerides and triglycerides) 2. Esters of long chain monohydric alcohols (commonly called “waxes”) 3. Sterol esters (sterol plus fatty acid) 4. Glycerol ethers (containing palmityl, stearyl or oleyl alcohols) B. Compound Lipids (esters of fatty acids and alcohol containing one or more additional groups) 1. Phosphoglycerides (commonly called phospholipids; esters of phosphate and free a hydroxyl of an a, 8, diglyceride or monoglyceride) (a) phosphate-monoester (phosphatidic acid) (b) phosphate-diester (e.g. phosphatidylcholine, phosphatidylethanolamine, phosphatidylserine) (c) phosphate-triester (complexes of phosphatidylethanolamine or phosphatidylserine) 2. Sphingolipids (sphingosine containing N-fatty acyl group) (a) sphingomyelin (containing phosphorus and choline) (b) cerebrosides (containing sugar; usually galactose) (c) sulfatides (cerebroside plus sulfate residue) (d) gangliosides (commonly called mucolipid; contains sialic acid) C. Derived Lipids (products of A. and B. still possessing lipid characteristics) 1. Fatty acids 2. Long-chain alcohols 3. Sterols 4. Terpenes 5. Hydrocarbons 6. Sphingosine D. Complex Lipids 1. Lipoprotein 2. Proteolipid (lipid plus peptide) 3. Phosphatide peptide (contains sphingosine, inositol, phosphorus and nitrogen)

leads to profound changes in the nature of the lipid (e.g. oxidation) and the lipid becomes virtually useless for further analysis. It is therefore not unusual to express the lipid content in terms of fresh or wet weight. This procedure can lead to large errors due to the fluctuations in the

LIPID METABO LISM A N D F U N C T I O N I N I N S E C T S

73

water content of the insect. From our experience, lean dry weight is a more constant value and is obtained by extracting the tissue with organic solvents (ethanol-ether or chloroform-methanol) and weighing the air-dried residue. The lipid can then be treated gently (e.g. evaporation under nitrogen) and used further for identification of its individual components. The lean dry weight then is the dry weight minus the weight of the lipid. For reviews of lipid extraction procedures see Haahti (1961) and Horning (1964). As Table I1 demonstrates, there is a wide variation in lipid content of insects of different orders and even within a single family. This is not difEcult to understand when one considers the large diversity of insects, TABLE I1 Lipid content of various insect species ~~

Insect COLEOPTERA Bruchidae Pachymerus bactris Callosobruchus chinensis Laria irresecta Buprestidae Eurythyrea marginata Carabidae Pterostichus vulgaris Pterostichus nigrita Pseudophonus pubescens Harpalus pubescens Harpalus griseus Cebrionidae Cebrio gigas Cerambycidae Ergates faber Rhagium inquisitor Plagionotus arcuatus Cerambyx scopolii Chrysomelidae Leptinotarsa decemlineata ¶¶ ¶¶

,,

¶¶ ¶¶ ¶¶

Galerucella luteola Colaspidema atrum

Stage

Per cent lipid

41 D 33-2W 23.6W 12.2w

57w 4.8W 28*4D 4.8W 6.0W 144W 13.0W 7.0W 14.4W 8.5W 4.2W 3.4w 13.4W 39.6D 7.0W 3.9w continued

74

LAWRENCE I . GILBERT

Insect Aulacophora fumolaris Cistelidae Heliotaurus menticornis Curculionidae Balaninus elephas Rhynchophoruspalmarum Anthonomus grandis

,,

Dermestidae Dermestes sp

,,

9,

Dytiscidae Dytiscus marginalis Lampyridae Luciola vitticollis Meloidae Mylabris pustulata Lytta vesicatoria Scarabaeidae Oryctes nasicornis Melolontha hippocastani Melolontha vulgaris Cetonia aurata Phyllophaga rugosa Geotrupes stercoralis Popillia japonica 99

9, 99

9,

9,

,,

Tenebrionidae Tenebrio molitor 9,

,,

9,

3,

,,

,?

LEPIDOPTER A Arctiidae Arctia caja Estigmene acraea Bombycidae Bombyx mori ,¶

99

,, 9,

Stage

Per cent lipid 22

w

6 W 28 W 22.3W 12.8W 35 D 47 D 21 D 6.2W 7.3w 4.8W 12.5D 13-5D 15.0W 6.1W 16.9W 1.9W 9.0 7.7 4.0W 3.2W 3.8W 4.0W 3.8W 17.4W 154W 14.9W 12.9W

4.1W 13.OD 12 w 343W 24.4D continued

75

LIPID METABOLISM AND FUNCTION IN INSECTS

Insect Bombyx mori 99

Carposinidae Carposina niponensis

,,

,,

9)

3,

9,

,,

Citheroniidae Citheronia regalis Eacles imperiaris

27.6W

Gelechiidae Pectinophora gossypiella

,, ,, ,, ,,

99

,, ,,

9,

Hesperidae Acentrocneme hesperiaris Lasiocampidae Malacosoma franconicrim Malacosoma americanum

,,

99

Lithosiidae Asura conferta Lymantridae Lymantria dispar Euproctis chrysorrhoea (Linn.) Noctuidae Alabama argillacea Heliothis armigera Prodenia ornithogalli Laphygma frugiperda Lycophotia margaritosa Euxoa segetum Agrotis segetum 99

¶3

9)

9)

27*1W 85.4D 35-7w 81.6D 17.4W 55W 6*0W

Cossidae Cossus ligniperda Danaidae Danaus plexippus

99

Per cent lipid 5.9w 9.1W

99

99

Stage

Laphygma exempta solitary

23 W 12 w 15.3W 44.8D 16.7W 50.5D 16-1W 33*4D 10

w

1.9w 82 D 24.7D 1.ow 1.5w 4-9w 24.4D 20.9D 18.1D 13.1D 0.9W 6.4W 23.1D 30.1D 13.5W 15W continued

76

LAWRENCE I . GILBERT

Insect Laphygma exempta gregarious Spodoptera abyssinia solitary Spodoptera abyssinia gregarious Spodoptera abyssinia solitary Spodoptera abyssinia gregarious Acronycta rumicis Colocasia coryli Daseochaeta at'pium Scoliopteryx libatrix PIusia gamma Notodontidae Pha lera bucephala Nymphalidae Vanessa urticae Olethreutidae Carpocapsa pomenella Papilionidae Papilio troilus Papilio turnus Papilio zolicaon Pieridae Pieris brassicae 99

>¶

Pieris napi 9)

Y9

Pieris rapae 9,

99

,,

,,

9,

99

9,

$9

,I

Y9

Psychidae Thyridopteryx sp. Pyralidae Galleria mellonella 99

>,

Myelobia smerintha Ephestia figulilella Loxostege similalis Loxostege sticticalis

Stage

Per cent lipid 2.7W 3.1W 4.9w 7.1 W 9.3w 25.5D 29.5D 12.5D 18.2W 334W 34.8W 10.3W 3.7w 44.2D 3-3w 4.6W 7.6W 10.8W 1.9w 6.0W 2.2w 3.6W 16.0d 7.7D 2.3W 12.1D 6.4W 26.1D 11.4W 29.7D 5-6W 9.1w 56-2D 21-0w 22 w 21-6W 18.8D 29.7D

continued

LIPID METABOLISM A N D FUNCTION I N INSECTS

Insect Loxostege sticticalis

,,

Y9

,,

9)

,,

9,

Pyrausta nubilalis 3,

YY

99

9,

Stage

77

Per cent lipid 5-9w 7.5w 30.2D 96W 3.2W 26.5W 9-7w 143W

Saturniidae Saturnia pyri Antheraea pernyi Y9

9,

Y9

,,

*Antheraea mylitta *Antheraea polyphemus *Antheraea roylei *Hyalophora cecropia *Hyalophora euryalus *Actias luna Adelocephala heiligbrodtii *Attacus atlas

*Attacus canningii *Samia Cynthia Automeris io

* Callosamia promethea * Calosaturnia mendocino *Rothschildiaforbesi

*Rothschildia orizaba

4.0W 1.ow 4*6W 14.8D 17.8D 4-OW 6.0W 21.6W 6-1W 38.3W 5.ow 216W 4-2W 22.8W 4.4w 17.6W 1*9W 2.8W 9-9w 41.3W 8.3W 40-OW 8.0W 36-7W 9.8W 18.2W 2.9W 13.4W 2.9W 22.8W 294W 3.7w 32.2W

Sphingidae Deilephila sp. Sphinx elpenor

35w 4.9w continued

78

LAWRENCE I . GILBERT

Insect

Stage

Sphinx ebenor Sphinx ligustri

,, 39

6-1W 7.8W 3.8W 6.1W 2*8W

9,

9,

Sphinx euphorbiae Thaumetopoeidae Thaumetopoea pityocampa

35w

HEMIPTERA Aphididae Aphis rumicis Aphis rosae Aphis fabae Hyalopterus pruni Coccidae Coccus cacti Eriosomatidae Pemphigus utricularias Jassidae Euttetix tenellus

7.5 w 6.5W 12 w 6.2W 8 W

20

9,

,,

9,

36.3D 11-6W 5-2W 6-2W 6-2W 32.7D

ORTHOPTERA Acrididae Acrida bicolor Anacridium aegyptium Orthocanthacris aegyptium Dociostaurus maroccanus Oxya japonica Locusta migratoria 9s

,,

Locusta migratoria solitaria

iAi ,, Locusta migratoria migratorioides ( A ) (A) Locusta migratoria solitaria (N) 9)

99

99

99

w

34.2D 11.5D

Lygaeidae Blissus leucopterus Oncopeltus fasciatus Notonectidae Notonecta glauca Pyrrhocoridae Pyrrhocoris apterus 99

Per cent lipid

,

9,

I

3.2W 3-8W 2.6W 3.3w 3 w 24W 2.8W 2.8W ll*ID 3-6W 14.0D 7.5D continued

79

L I P I D METABOLISM A N D F U N C T I O N I N INSECTS

Insect

Stage

Locusta migratoria migratorioides

(N) ?(A) $.(A) (A) (A) (A) (A) (A)

,>

99

Locusta pardalina solitary Locusta pardalina solitary Locusta pardalina gregarious 9,

,,

9,

Per cent lipid 10-7D 10.4W 147W 2.9W 12.8D 3.7w 14-6D 10 w 5.3D 9.3D 3.8W 3-2W 12 w 3.5w 25w

Schistocerca gregaria Schistocerca gregaria solitary (N) Schistocerca gregaria gregarious (N) Pezotettix giornae ?(A) Melanoplus atlanis (A) Melanoplus differentialis (E) 99 (A) Sphenarium purpurescens (A) Bacunculidae Dixippus morosus (A) Blattidae BIatta orientalis (A) ,* (A) BlattelIa germanica (N) ,, ?(A) d(A) (A> Periplaneta americana ?(A) $.(A) (N) ?(A) $.(A) Tettigoniidae Tettigonia uiridisima (A) 9,

2*4W 4.3w 3.2W 5.7w 4.8W 1-7W 17.1D 13.9D 14.5D 7.7w 8.9W 7-1W

9,

9

9,

9,

9,

73

3.8W

ODONATA Aeschnidae Aeschna sp. ISOPTERA (unspecified)

25w

(N)

6.1W

(L)

5-6W

(L) ?(A) $.(A)

30-OD 3.3w 34w

DIPTERA Anthomyiidae Pegomyia ulmaria Calliphoridae Calliphora erythrocephala

,,

')

continued

80

LAWRENCE I . GILBERT

Insect Calliphora erythrocephala Calliphora vomitoria Lucilia sericata

,.

9,

99

9,

,,

99

Phormia terraenovae Phormia regina 99

3,

9,

,,

Chironomidae Chironomus sp. Tanytarsus lewisi Culicidae Culex pipiens 3,

9,

Y9

99

Anopheles atroparuus Anopheles stephensi Anopheles gambiae

Glossinidae Glossina palpalis Glossina morsitans

Muscidae Musca domestica

Oestridae Gastrophilus intestinalis Syrphidae Eristalis tenax

Stage

Per cent lipid 12.7D 7.8D 19 D 31 D 27 D 20 D 12.2w 8.9W 6.8W 52W 8.3W 11.7D 27.9W 3-7w 32*7D 4.0W 5-9w 57w 8.4W 15-OD 9.8D 22.7D 21.9D 16.8D 352? 2.7W 7.1W 6-3W 4.2W 2.5W 4.9w 4.2W 2.6W 2*9W

5w 1.8W continued

LIPID METABOLISM A N D F U N C T I O N I N INSECTS

Insect

Stage

81

Per cent lipid

HYMENOPTERA Apidae Apis dorsata Apis melrifera workers

,, ,, ,, 9,

,9

,, ,9

,,

9,

,, ,, ,, ,, ,, ,, ,, ,,

7,

),

,, 77

,, 9,

Y,

,, 9,

,, drones 79

freshly hatched wax producing forager winter worker Queen worker Queen summer winter

Formicidae Camponotus vagus Cremastogaster scuiellaris Tenthredinidae Croesus septentrionalis Vespidae Vespa cincta

(N) (P) (A) (PI (A) (A) (A) (A) (A) (L) (L) (P) (P) (A) (A)

4.7w 4.1W 0.9w 6.1W 1.5W 9.4D 10.2D 95D 7.3D 3-7w 4.9w 3.7w 5-3w 13.OD 12*2D

(L)

(A)

2.4W 10.9W

(PP)

26.OD

(I-)

67W

(E)egg; (L)larva; (N)nymph; (PP)prepupa; (P)pupa; (A) adult; (W) wet weight; (D) dry weight; * abdomens only. Modified from review by Fast (1964); consult Fast (1964) for original references and other data.

their modes of living and varying habitats. In most insects the female contains more lipid than the male, as lipid is a most efficient substrate for egg development. However, the reverse may be true for many species and this is especially evident when we consider the Lepidoptera (Table 11). This must be borne in mind since most of the author's research has utilized the American silkmoth Hyalophora cecropia, and this insect w ill be used to exemplify many points in this review. B . ALTERATIONS D U R I N G METAMORPHOSIS

In the course of a study on the alterations in juvenile hormone content during the life cycle of H.cecropia, we noted a sexual dimorphism in lipid content in the adult stage, and conducted a number of experiments to determine its basis (Gilbert and Schneiderman, 1961b). By preparing lipid extracts of insects at various stages in the life cycle the observations in Table I11 were made. Lipid content per individual larva

00 h)

TABLE I11 Lipid content of H. cecropia from egg to adult emergence

Stage

Number of animals

unfertilized eggs embryos 1st instar larvae 2nd instar larvae 3rd instar larvae 4th instar larvae 4th instar larvae 5th instar larvae 5th instar larvae 5th instar larvae prepupae Diapausing pupae 3 Diapausing pupae 9 Chilled pupae d Chilled pupae d Chilled pupae 0 Chilled pupae 0 Developing adult 3

750 1000 200 20 17 6 6 1 4 3 3 20 20 20 2 20 2 4

Time sacrificed freshly laid 7 clays

Fresh weight (€9

Lipid Lipid Dry Percent Grams as % as % weight dry lipid: fresh dry (g) weight individual weight weight

3.80 4.30

0.00025 0.00019

4.94 4.30

0.00012 0.00109 0.00922 0.0204 0-0416 0-0460 0-1189 0.2538 0-3723 0.2890 0.3150 0.3970 0.27 18 0.2435 0.2135 0.3100

4-53 2.27 0-98 1.34 1-28 1.40 1.63 1.86 5-68 7-34 5.57 8.68 6.44 5.43 4-24 8.67

0.516 newly hatched 0.957 early 15.879 late 9.122 early 19.463 late 3.285 early 29.233 mid 40-976 late shortened body 19.6648 1 month old 78.8 113.1 1 month old 6O, 6 months 9 1-50 6O, 6 months 8.439 6O, 6 months 91.50 6O, 6 months 10.067 day 2 14.30

0.125 0-131 2.316 1~207 2-300 0.334 5.141 6-690 5.361

24.22 13-69 14.59 13-23 11.82 10.17 17.59 16-33 27.26

-

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2.209

26.19

2-485

24.69

-

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Comment

includes chorion includes chorion and yolk 18.72 whole animal 16.56 whole animal 6.77 whole animal 10.13 whole animal 10.85 whole animal 13.77 whole animal 9-25 whole animal 11.38 whole animal 20.84 whole animal - whole animal whole animal whole animal 24-59 whole animal whole animal 17.18 whole animal - abdomens only and pupal cuticle

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LIPID METABOLISM A N D FUNCTION I N INSECTS

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84

LAWRENCE I . GILBERT

was observed to increase with age and size as expected, but the rate of increase varied in the various instars when compared to the net increase in non-lipid constituents. The newly hatched larva contains an appreciable amount of lipid, but this percentage decreases to a more or less constant value in the fourth and fifth instars (1-3-1.9% fresh weight; 9-3 to 13.8% dry weight), although the lipid content increases as the larva grows. The prepupa loses water rapidly and lipid content increases, so that at pupation lipids constitute between 5 and 7% of the wet weight of the insect. At this time a sexual dimorphism in lipid content is evident; the male pupa contains 50% more lipid than the female per gram fresh weight. This dimorphism also exists in pupae chilled for 6 months at 6". During the pupal-adult transformation this dimorphism increases markedly until in the adult moth the tissues of males contain about 5 times as much lipid per gram fresh weight as the tissues of the female. During adult development, the female apparently utilizes far more lipid than the male (see also Domroese and Gilbert, 1964). A 5 g previously chilled male pupa contains about 360 mg of lipid while the newly emerged adult moth contains approximately the same amount. For the female, the picture is quite different. A similar 5 g female pupa contains about 230 mg of lipid. The correspondingnewly emerged adult with eggs contains about 185 mg. In short, during adult development females appear to use more lipid than males. Because the abdominal tissues of female moths have a greater water content than those of males, the sexual dimorphism in lipid content is not as impressive when calculated on a dry weight basis. The results of a typical series of extractions reveal that during the first week of adult life the abdomens of males contain from 51-79% lipid per gram dry weight while the abdomens of females contain from 19-25%. After the sixth day of adult life there appears to be a rapid utilization of lipid in the males indicating a significant increase in catabolism at this time. The same is true of female moths, with the most rapid utilization occurring after the tenth day of adult life: senile females may contain as little as 8% lipid per gram dry weight as contrasted with about 20% in young females. To determine the extent of the lipid metabolism associated with egg development during adult development, a series of female pupae were , castrated and allowed to develop. Two days after eclosion the moths were extracted and the lipid content determined. The abdomens of castrated females contained about twice as much lipid as normal eggless female abdomens but still less than as much as male abdomens (less

+

LIPID METABOLISM A N D FUNCTION I N INSECTS

85

than 3% on a fresh weight basis). Thus, although egg development involves some utilization of lipids, it cannot by itself account for the sexual difference in lipid content between males and females. It also appears that during adult development both normal and castrated females utilize far more lipid than males. Castration of males had no effect on their lipid content. When ovaries were implanted into male pupae, the adults which emerged contained large masses of eggs. Extraction revealed that these egg-filled male abdomens contained less lipid than normal abdomens but twice the amount of those of castrated or normal females. This result implies that egg development involves lipid catabolism but that certain non-ovarian tissues of the developing female utilize more lipid than corresponding male tissues. Parabiotic union of male and female pupae resulted in parabiotic moths. This technique resulted in the elevation of the lipid content of the female but a normal content in the male. What conclusions can we draw from this data? As was expected, the lipid content of individual larvae increased with size and age reflecting the synthesis of lipid from leaf material. Since no attempt was made to sex the larvae, nothing can be said regarding any sexual dimorphism in lipid content at this stage. The 50% decrease in lipid content per individual between egg and first instar larva indicates lipid utilization during embryogenesis. Although we shall return to this point later, this does confirm observations made on insects of other orders (Rothstein, 1952)and on B. mori (Niemierko et al., 1956), although the lipid content is much lower in H. cecropia eggs (as per cent fresh weight) than in the diapausing egg of Bombyx. This could be due to both the greater weight of the chorion of the Cecropia egg and to greater substrate storage in the diapause egg which supports a living embryo over winter. In the fifth instar larva when the water content reaches the extraordinarily high level of about 90% of the total weight, the animal begins to synthesize lipid at a high rate. The fifth instar larva has about five times as much total lipid as the early fifth and a lower water content. During spinning, when a large quantity of protein is used for constructing the cocoon, there is a further increase in total lipid and a loss of water. Whether this large increase in lipid during spinning reflects a release of lipids firmly bound to the proteins used in construction of the cocoon or whether the increase is a result of enhanced lipid synthesis has not been definitely answered. The fact that male pupae and adults contain more lipid than females seems peculiar to Lepidoptera. In most insects, the female usually contains more lipid (Table 11). Our results are consistent with the findings of

86

L A W R E N C E I . GILBERT

Niemierko et al., (1956; see also Vaney and Maignon, 1906; TimonDavid, 1930; Yamafuji, 1937) in Bombyx mori, and Demyanovsky and Zubova (1957) in Antheraea pernyi. Niemierko and his colleagues found that the lipid content of newly emerged male Bombyx was 48% per gram dry weight while that of the female was only 25% (17.5% in males and 6% in females on a fresh weight basis). The females used 50% of their stored lipid between spinning and adult emergence while the males utilized only 30%. In A . pernyi, Demyanovsky and Zubova found the sexual dimorphism in lipid content evident as early as the fifth instar larva and it continued until adult emergence. Our results (Gilbert and Schneiderman, 1961b) extend this dimorphism to a total of seven families of Lepidoptera. The above results indicate that lipid is used during the pupal-adult transformation of female Lepidoptera but that if any lipid is used for energy to underwrite the many syntheses occurring during construction of the male adult, other substrates are presumably being converted to lipid. Biochemical systems that accomplish such conversions have already been established (see Section IV). Why is there a lipid-sparing system operating during adult development of male Cecropia? The higher concentration of lipid in the male moth is most likely correlated with mating behaviour, since the male moth flies relatively great distances in search of the virgin female. As we shall see subsequently (Section 111) the male appears to utilize lipid as the primary substrate for flight. The monarch butterfly is a continental migrant and also utilizes lipid during flight. Prior to migration this specieslays down a store of fat as do migratory birds (Beall, 1948). Since its only food is nectar which does not contain lipid, it evidently converts carbohydrates to lipid during periods of repose and then uses it during flight. In migratory moths, it appears that there is no sexual dimorphism in lipid content (Williams, 1945) since both sexes migrate and presumably use equal amounts of lipid. The fact that the female Cecropia moth utilizes a significant portion of her pupal lipid during adult development and contains much less than the male, can be explained in part by her adult role. First, the female converts a large percentage of her endogenous substrate into the egg (yolk, cytoplasm, chorion, etc.). Secondly, the female does only limited flying after emergence and therefore needs little flight fuel. A by-product of the female’s lipid utilization during adult development is metabolic water, which is reflected in the greater water content of the female as compared to the male. During adult life, lipid appears to be the favored substrate of both

L I P I D METABOLISM A N D F U N C T I O N I N INSECTS

87

sexes. The utilization of lipid at this stage when the animal has left its desiccation-proofcocoon and heavy pupal cuticle, provides these moths with enough metabolic water to offset desiccation. It is possible that the male converts protein to lipid during the first week of adult life when the lipid content drops only slightly, since there is very little carbohydrate present at this time (Domroese and Gilbert, 1964). Perhaps these non-feeding adults ultimately die of lipid depletion. What is the basis of the sexual dimorphism in lipid content in these Lepidoptera? Our data reveal that egg development can explain in part

0.72

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FIG. 1. The respiratory quotient of male H. cecropia during adult development and adult life. Determinations were made on five animals, each designated by a different symbol. (From Domroese and Gilbert, 1964.)

the low lipid content of female moths. However, the surgical operations performed never reversed or even equalized the lipid differences between the sexes. Since this dimorphism exists in the pupal and even in the larval stages (Demyanovsky and Zubova, 1957) we believe it to be a genetic difference which may find its first phenotypic expression during embryonic life. The above experimental findings were based almost entirely on direct lipid extraction of animals at different stages of development. To explore this sexual dimorphism further, we (Domroese and Gilbert, 1964) determined the respiratory quotients (RQ) of both sexes of H . cecropia during pupal-adult development. The results of these determinations show a differencein pattern between males and females (Figs. 1,2). In the female, kA.1.P.

4

88

LAWRENCE I . GILBERT

the RQ decreases from higher values early in development to values between 0.75-0.80 shortly after the first week of development. The RQ remains at this characteristic level throughout the rest of development and in adult life. In terms of the role of lipid as an energy source for development, the RQ pattern suggests that after the first week of development lipid provides a greater portion of the energy expended in the female. In the male, the RQ is maintained for the most part at a level well above 0.80 until the seventeenth day of development, at which time it falls to levels indicative of greater lipid utilization. The RQ persists at this lower 1.00

8

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FIG.2. The respiratory quotient of female H. cecropia during adult development and adult life. Determinations were made on five animals, each designated by a different symbol. (From Domroese and Gilbert, 1964.)

value during the remainder of adult development and in adult life. This drop in RQ characteristically occurred simultaneously with the appearance of the large dark pigmented wing spot on the seventeenth day of development. This RQ pattern as well as the data previously cited indicates that the male uses more of the non-lipid substrates for energy during adult development with a conservation of the lipid stores for use during adult life. In Section 111, we will discuss in detail the role of lipid in the adult life of the male moth. Among the various deductions that can be made regarding the above/ one fact is quite clear. The amount of lipid varies considerably during the life of an insect as well as between various species. The difference in the quantity of lipid expressed as per cent fresh weight has been reported to

L I P I D METABOLISM A N D F U N C T I O N I N INSECTS

89

vary from 1% in some Lepidoptera to 50% in the beetle Puchyrnerus (Niemierko, 1959). Examples of the high lipid content of some insects can be seen in the older work of Maciuca (1935) who showed that in the fall, Pyrrhocoris upterus contains 32.7% lipid (dry weight) while Fulton and Chamberlain (1934) noted that the lipid content of E. tenellus may rise as high as 42.2% of the dry weight of the female. Sinoda and Kurata (1932) showed that ether extracts of Dermestes larvae account for as much as 47% of the dry weight of the insect. As Scoggin and Tauber (1950) point out, many factors influence the lipid content of insects including stage of development, nutrition, environmental temperature, sex, starvation, diapause, cold hardiness, whether migratory or not and finally the systematic position of the organism under study. Rather than pursuing the most laborious task of discussing the lipid content of diverse insect species, let us now turn to the qualitative nature of insect lipids. C . NATURE OF INSECT LIPIDS

1. Glycerides In general, almost all of the compounds listed in Table I have been identified in one insect or another. These will be discussed in some detail throughout the paper, but there is no doubt that the major lipid component in insects as in other animals, is triglyceride (TGL) : the glycerol esters of long chain fatty acids. We can think of triglycerides as a secondary energy source since they must be hydrolyzed before energy can be made available to the cell. This results when the individual fatty acids cleaved from the glycerol moiety undergo B-oxidation within the mitochondria (see Section 111). In general, these TGL form the greater part of the lipid content of the insect at all developmental stages (Fig. 3). In Muciosiphium burri, about 80% of the extractable lipid is TGL and less than 3% exists as unesterilied fatty acid (FFA) (Strong, 1963a, b). This predominance of neutral lipid has been demonstrated in many insects, both in total body extracts (cf. Fast, 1964) and isolated fat body (Chino and Gilbert, 1964, 1965a; see also reviews of Kilby, 1963 and Gilby, 1965). Large quantities of FFA have been reported in the older literature (cf. Scoggin and Tauber, 19Sl) but in many cases one can “ see” the esterified lipids hydrolyzing before one’s eyes while reading the methods sections of these papers (see, however, Albrecht, 1961). In a study of the lipid of adult Anthonornus grundis by silicic acid chromatography, Lambremont et ul. (1964) showed that newly emerged

90

LAWRENCE I . G I L B E R T

adult boll weevils had 2 4 % body lipid of which only 2% was triglyceride. After 2-3 weeks of feeding, non-diapausing adults contained 6 1 0 % lipid (40-60% of which is TGL), whereas diapausing adults had 18-25% body lipid containing 75-85% TGL. It thus appears that lipid serves as the energy source for this diapausing insect which must last over winter, a suggestion made earlier by Braze1 and Newsom (1959).

Tube no.

FIG.3. Column chromatography of lipids from male H. cecropia moths. The column support was Florisil and elution was according to Chino and Gilbert (1965a). Tubes 12 to 22 represent triglycerides. (From Domroese and Gilbert, unpublished observations.)

Lambremont and his colleagues also demonstrated that the major fatty acids in the boll weevil were oleic and palmitic acids, and that the diet of the adult was the determining factor controlling the type of fatty acids incorporated into TGL.

2. Fatty acids (a) General characteristics. Protoplasmicfatty acids usually vary from C2to C,,in chain length and are most commonly composed of an even number of carbons, with oleic acid being the most abundant in nature. Of the many unsaturated fatty acids, the majority have a double bond between Cgand Clo (Fig. 4). Isomers of any particular fatty acid differ from one another by the position of the double bond, cis-trans isomerism, or both. There is a large variety of fatty acid types including

LIPID METABOLISM A N D FUNCTION I N INSECTS

91

branched, unsaturated (up to six double bonds), hydroxy saturated or unsaturated, and cyclic. Saturated fatty acids having ten or less carbon atoms are liquid at room temperature and are thus easier to handle in experimental situations although more prone to auto-oxidation. All fatty acids with the exception of acetic acid have a density of less than one. The most common form for fatty acids in living things is as constituents of glycerides which are usually mixed (i.e. contain more than one variety of fatty acid per molecule). Although we speak of unesterified fatty acids as free fatty acids, in the cell or in the hemolymph they are usually in the form of the cationic salt (e.g. sodium palmitate) and can be referred to as palmitate or oleate rather than palmitic acid or oleic acid. 0

1I

CH3(CH2)7CHaCH2(CH&C--OH I. Stearic acid

0

It

CH~(CH&CHL-CN(CH~)~C-OH 11. Oleic acid (AS:l0) 0

CH3(CHa)rCHSHCHaCH=CH(CH& !! -OH 111. Linoleic acid (Aa:loJ2:13 )

i

CH3CH2CHSHCHaCH=CHCH2CH=CH(CH2)7 -OH IV. Linolenic acid (A9:10.12:13.15:16 1

FIG.4. The structure of some important C,, fatty acids.

(b) Fatty acid composition. The fatty acid composition of numerous insect species has been reviewed by Fast (1S64), and only the more recent work will be discussed here. The older literature suffers from a lack of sophisticated analytical tools (i.e. gas chromatography) and in many cases the unsaturated fatty acids were handled without respect, undoubtedly leading to oxidative deterioration in many cases. In a study of the lipids of homopterous insects, Strong (1963a) determined the fatty acid composition of 21 species of aphids and 6 species of leafhoppers. The percentage of the various fatty acids varied quite a bit between groups and even between species of a single group. In general, the aphids had a relatively large percentage of C14saturated fatty acid while the leafhoppers had a predominance of C18(especially

92

L A W R E N C E I . GILBERT

C,,,,) fatty acid. Strong concludes that “the fatty acid composition of an aphid appears to be a species characteristic not especially influenced by the host plant.” Honeydew secreted by Myzus persicae contains for the most part free sterols and fatty acids and appears to originate from the phloem sap (Strong, 1964). It seems from Strong’s work that the fatty acid composition of a species is rather specific and if this is true, it may be that gas chromatography is the “ultimate weapon” for the taxonomist. In examining the fatty acid content of the boll weevil, Lambremont and Blum (1963) identified 23 fatty acids varying in chain length from six to twenty carbons. Of these, 8 fatty acids (myristate, palmitate, palmitoleate, heptadecanoate, stearate, oleate, linoleate, linolenate) constitute 98% of the total, with 62% of the fatty acids containing at least one point of unsaturation. Palmitate makes up about 31% of the total while oleate constitutes about 30% of the total. Of the eight predominant fatty acids in the boll weevil, all but palmitoleate, heptadecanoate and stearate were identified in Drosophila melanogaster (Anders, 1960). It is possible that Anders’ paper chromatographic system was too insensitive for detection of the other three. Fawzi et al. (1961) have also shown that palmitate and oleate are the most common fatty acids in the TGL of Locusta migratoria. Almost all of the studies on fatty acids in insects utilized extraction of whole insects and generally palmitate and oleate predominate. The situation in the arachnids seems to be similar (Blum et al., 1963). Other more exotic fatty acids have been identified in insects and insect products, but are the exception rather than the rule. In bees for example, hydroxy fatty acids are common in queen bee larvae and royal jelly. Royal jelly is of course the nutrient fed to young larvae causing them to develop into queens. In 1957, Butenandt and Rembold identified 10-(OH)-decenoic acid in royal jelly. Weaver and Law (1960) studied the heterogeneity of fatty acids in this most interesting nutrient and showed the presence of: decandioic acid; monohydroxymonocarboxylic acids ; dihydroxymonocarboxylic acids. Weaver et al. (1964) reported that 9-13% of the dry weight of royal jelly is lipid of which 90% is FFA. Pain and her colleagues (1962) had demonstrated previously that royal jelly contains adipic, pimelic and suberic acids. Brown and his group (Brown and Felauer, 1961; Brown et al., 1961) showed the presence of 9-(OH)-dec-Zenoic acid, lO-(OH)-dec-2-enoic acid, 10-(OH)-decanoic acid, dec-Zendioic acid, p-hydroxybenzoic acid and sebacic acid. The presence of these acids is now known but their function is still a matter of speculation. They may have an anti-

LIPID METABOLISM A N D FUNCTION I N INSECTS

93

bacterial action (cf. Slepecky and Gilbert, 1962) or may have morphogenetic effects in inducing those larvae fed on royal jelly to develop into queens. Several of these fatty acids and their derivatives have been identified in the products of hydrolysis of queen bee larval lipid (Pain et al., 1962). Identified by the French group in larval lipids were myristate, palmitate, dec-Zenedioic acid, suberic acid and trans-10(OH)-dec-Zenoic acid. This latter acid accounted for about 20% of the total fatty acids present but its function is unknown. The wax of Apis mellifera was thought to be composed largely of fatty acids and alcohols containing the even numbered homologues of CZ4to C34fatty acids (Chibnall et al., 1934) but the use of gas chromatography has revealed the presence of some odd numbered fatty acids as well (Downing et al., 1961). The cuticular wax of B. mori is composed for the most part of fatty acid-alcohol esters with fatty acid chain lengths of from c16 to C,, (Amin, 1960; Bergmann, 1934; Shikata, 1960). The most complete survey by one individual of the fatty acid constitution of insects is that of Barlow (1964), who by gas chromatography analysed 30 species of insects (Table IV). Unfortunately, only one analysis was conducted on most of the species due to lack of material. Barlow’s findings indicate that the fatty composition is characteristic of the species. For the family aphidoidea (Homoptera) for instance, more than SO% of the fatty acid was saturated C14 whereas in most species outside of this group, this fatty acid only constituted about 15% (19 to 60%) of the total. All the Diptera had a high proportion of while most other species yielded no more than about 2.2% of this acid. That some insects can synthesize this c16:1is no longer in doubt since A . aflnis contained a high concentration of this fatty acid even when grown on a diet devoid of it. In general, Barlow’s work suggested that the stage of the insect had little effect on the fatty acid composition (see, however, the subsequent discussion of the work of Herodek and Farkas). [The new rash of papers on fatty acid composition of insects (and other living things) is due in great part to the classic efforts of Martin and Synge (1941) who first described the concept of gas chromatography in which gas is the transient phase rather than a liquid (as in paper or column chromatography).] A relationship appears to exist between the temperature to which a tissue is constantly exposed and the degree of unsaturation of the fatty acids present in the tissue. This appears to be true of all animals and has been demonstrated in insects. Newly emerged female Culex tarsalis for example possess a greater percentage of unsaturated fatty acids

TABLE IV Total fatty acid composition of some insect lipid

% fatty acid1 Stage

Insect Coleoptera Calosoma calidum (Fabricius) Harpalus caliginosus (Fabricius) Sitona scissifrons (Say) Leptinotarsa decemlineata (Say) Trirhabda virgata Lec. Pyropyga decipiens Harris Tetraopes tetraophthalmus (Forster) Neuroptera Corydalus cornictus (Linnaeus) Trichoptera Unidentified Homoptera-Aphididae Dactynotus ambrosiae (Thomas) S.L. Aphis pomi de Gier Acyrthosiphon pisum (Harris) Tuberolachnus salignus (Gmelin) Pemphigus populicaulis (Fitch) Homoptera-o ther Cicadidae unidentified Campylenchia latipes (Say) Stictocephala diceros (Say) Philaenus spumarius (Linnaeus)

12

14

16:O

16:l

18 13 10 11 9 11 8

5

2 3 1 1 1 2

adult adult adult Pupa adult adult adult

<1

larva

21

10

22

11

larva

1

1

46

10

nymphs and adults nymphs and adults nymphs and adults nymphs and adults nymphs and adults

1 1 9 4 3

9 9 4 3 3 5 5 7 8 5 3 2

3 4 3 4

adult adult adult adult

-

1 10 25 3

2 2 1 1 1 2 2 1

1 1 99+1 1 12 1

5

-

3 3 3 1 1

18:O 18:l 2 2 10 10 13 10 1 4

30 23

18:2 18:3

25 16 42 6

21 58 12 32 25 9 16

6

14

8

5

35

1

50

21

-

14 19 25

2

12

c1

8

7

-

-

5 3 2 1 1 3 5 4 4

4

25

12 16 4 11 63

-

-

-

-

-

14 11

1

2

1

4

15 48

66 23

?

M

Diptera Agria afinis (FallCn) Musca domestica (Linnaeus) Drosophila melanogaster Meigen Aedes aegypti (Linnaeus) Hylemya brassicae (BouchC) Lepidoptera Nymphalis antiopu (Linnaeus) Galleria mellonella (Linnaeus) ffyphantria cunea (Drury) Calophasiu lunula (Hufnagel) Hymenoptera Neodiprion sertifer (Geoffroy) Acunthomyops claoiger (Roger) Acanthomyops claoiger (Roger) Orthoptera Melanoplus sanguinipes (Fabricius)

19 28 22 20

Pupa Pupa Pupa Pupa Pupa

1 1 2 1 1

2 4 20 3 4

Pupa Pupa Pupa larva

_

6

<1

9

-

1 1

31 24 18

larva worker pupae sexual pupae

1 2 98

2 4 1

9 29 1

1

1

11

adult

-

_

15

29 21 21 19 60 <1 1 4 2

4 1

1 6

-

13 25 16 16

-

I 6

11 43 41 24

11 17 5

-

4 4

-

33 54

-

45 1

-

1

4

19

20

1

6

7

32 19 18 35 20

-

15

Lauric (12:0), myristic (14:0), palmitic (16:0), palmitoleic (16:1), stearic (18:0), oleic (18:1), linoleic (18:2), and linolenic (18:3) acids are expressed as a percentage of their total. (-): not observed. (from Barlow, 1964)

96

LAWRENCE I . GILBERT

when raised at low temperature (Hanvood and Takata, 1965). In addition, photoperiod appears to play some role in this phenomenon since a short day also causes a greater degree of unsaturation. Since these and perhaps other environmental conditions affect the consistency of the fatty acids in some insects, it is imperative that workers describe the conditions under which their experimental animals were raised. In some insects, the body lipid may contain up to 70% of polyenoic fatty acids with linoleic and linolenic acids the most commonly encountered (cf. Schmidt, 1964). Of these, some tetraenes and pentaenes are present. In Formica polyctena females for instance, fatty acids with two to five double bonds have been detected (Schmidt, 1963). In an analysis of the lipids of pre-pupating larvae of Phalera bucephala (Lepidoptera), Schmidt and Osman (1962) showed the lipid to comprise 32.7% of the dry weight, most of which was glycerides. Of the fatty acids present, palmitate, oleate and linolenate exceeded the concentration of stearate and linoleate. In addition, trace quantities of a tetraenic fatty acid were present. A tetraenic fatty acid was also identified in the total body lipid of adult Locusta migratoria by Fawzi et al. (1961). This insect’s lipid contains predominantly oleate, linoleate and palmitate plus lesser quantities of myristate, stearate, linolenate and arachidonate. In Musca domestica, there is a general trend toward synthesis of fatty acids during early larval growth and subsequent lipid utilization during metamorphosis (Pearincott, 1960). Alterations in individual fatty acid content do occur during metamorphosis but are usually masked when the entire FFA fraction is studied. Individual analysis of the fatty acid content during metamorphosis is rare. Herodek and Farkas (1960) conducted such analyses of unsaturated fatty acids during the embryonic and post-embryonic development of Bombyx mori. The percentage of total unsaturated fatty acids declined only slightly from embryo to last instar larva but fell from 80 to 70% in the prepupa. During adult development, the percentage of unsaturated fatty acids increased in the female where they may play a role in egg development, but fell in the male. On an individual fatty acid basis, the percentage of oleate, linoleate and linolenate remained rather constant during embryogenesis. Once the fist-instar larva hatched however, linoleate increased from about 10% of the total to more than 50% by the third instar, presumably at the expense of oleate and linolenate which decreased during this period. Subsequent to the third-instar, the concentration of linoleate decreased to reach a level of about 15% in prepupae after

L I P I D METABOLISM A N D F U N C T I O N I N INSECTS

97

which the concentration remained rather constant. Coincidentally with this decrease in percentage of linoleate, increases occurred in both linolenate and oleate. In all instances, the percentage of individual fatty acids was greater in the female imago than in the male. These results indicate that either there is an interconversion of linoleate to and from oleate and linolenate by hydrogenation and dehydrogenation, or a pronounced utilization of linoleate during the larval-pupal moult. Other than their value as a source of energy, little is known regarding the role of individual fatty acids in molting and metamorphosis. If the drastic alterations in fatty acid concentration discussed above can be extended to other species, it would appear that these unsaturated fatty acids may play a crucial role in the growth of the insect. With increasingly more sophisticated apparatus, future studies along this line are assured. According to Gilby and Cox (1963), the cockroach odor which is familiar to all of us who have worked with Periplaneta americana is due to an unidentified unsaturated fatty acid. Once we have a general idea of the fatty acid composition of insects there are several questions to be asked. How do these fatty acids come to be in the insect tissues and what is their purpose? Essentially then, we wish to know the biosynthetic scheme for fatty acids and their role in providing energy for the organism. Insects are famous (or infamous) for ingesting almost anything in the plant and animal kingdoms. When the food source is lipoidal or contains some lipid, degradation by digestive enzymes can lead ultimately to fatty acid. But how can an insect on a lipid-free diet build up a lipid store? To answer the last question we must entertain a discussion of molecular interconversion of carbohydrates and proteins to lipid.

111. LIPIDUTILIZATION A . DIGESTION A N D ABSORPTION

Since the bulk of ingested lipid is usually in the form of TGL which may not be able to penetrate the gut wall, the function of the digestive enzymes is to split these glycerol esters into simpler molecules, namely, diglycerides (DGL), monoglycerides (MGL) and FFA. We will not discuss these digestive processes in detail nor the transport of lipid from the digestive tract into the hemolymph because little is known regarding the former in insects and what is known regarding both phenomena has been recently reviewed (House, 1965; Treherne, 1962). The main

98

LAWRENCE I . GILBERT

digestive enzyme acting upon lipids is a lipase that mediates the hydrolysis of TGL. Contrary to what we might assume, hydrolysis most likely does not go to completion in the gut. In mammals for instance, a controversy has embroiled the field for many years regarding the exact form in which lipid passes from the gut to the circulatory system. In a concise review of the work in his laboratory, Isselbacher (1965) discusses the metabolism and transport of lipid by the mammalian intestinal mucosa. The major hydrolytic products of ingested lipid in the intestinal lumen are FFA and MGL. These end products then enter the mucosal cell where they are utilized in the resynthesis of TGL. The FFA are acylated and react with a-glycerophosphate to yield phosphatidic acid derivatives (i.e., glycerides and phospholipids, see Section IV). All the enzymes necessary for the synthesis of TGL appear to be located in the microsomal fraction of the mucosal cells. It may be of interest to those working on hormone action that mucosal microsomes from adrenalectomized rats have less than half the capacity of normal mucosal microsomes to esterify palmitate, indicating regulation of lipid absorption by the corticosteroids. In an excellent biochemical and electron microscopic investigation, Ashworth and Johnson (1963) found that the fatty acid released by the action of digestive lipase is taken into the intestinal epithelial cell by an infolding of the cell membrane. It is only when the fatty acid is internalized and transported further into the cell that it is resynthesized into TGL. It is then the TGL which is released into the intestinal lymphatic system. Studies of this calibre have not been reported in insects, but the need for such research is surely great if we are to understand the complete cycle of lipid utilization in insects. Digestive lipases have been the objects of study in several insects (Baker and Paretsky, 1958; Duspiva, 1934; Gaeta and Zappanico, 1959; Schlottke, 1937; Yamafuji and Yonezawa, 1935), and they appear to have the same function as the pancreatic lipase of mammals (cf. Desnuelle and Savary, 1963). In mammals, the main products of this hydrolysis are unesterified fatty acids and 2-monoglycerides, and there appear to be two competing reactions for the MGL absorbed into the intestinal cell. On the one hand there is the tendency to re-esterify to the higher glycerides utilizing acyl thioesters and on the other hand the tendency for the final hydrolysis of the MGL into FFA and glycerol (Senior, 1964). The major goal in the absorption of lipid in the mammalian intestine is the conversion of ingested TGL to lymph TGL with ’ the least expenditure of energy and minimal alteration in the structure of the fatty acids. In insects we know that the presence of FFA aids in the absorption of lipid out of the alimentary canal (Eisner, 1955; cf.

LI PI D METABOLISM A N D F U N C T I O N I N INSECTS

99

Treherne, 1962) but have little idea of the chemical form in which these materials enter the hemolymph. The aim of lipid absorption in insects is to convert ingested TGL into an unknown form of hemolymph lipid and thence to fat body TGL with a minimum expenditure of energy. The mechanics and biochemistry of this conversion are virgin research areas. In fact, there have been few detailed studies on the lipid content and fatty acid identification in the hemolymph. Wyatt’s (1961) excellent review on the biochemistry of hemolymph scarcely mentions lipids. In the recent 41-page review on the composition of insect hemolymph (Florkin and Jeuniaux, 1964) lipids are completely omitted. Hopf (1940) found that the larval hemolymph of Phormia contains about 1% lipid while Levenbook (1950) has demonstrated the presence of 137 mg % of fatty acids and sterols in the hemolymph of Gastrophilus larvae. More recently, Tietz-Devir (1963) has demonstrated the presence of 28.8 p equivalents of esterified fatty acids and 0.1-0.2 p equivalents of FFA in the hemolymph of L. rnigratoria. Most of the work in this connection deals with lipid transport from the fat body and will be discussed in Section 111. However, in one investigation of the hemolymph of the adult cricket, A . domesticus (Nowosielski and Patton, 1965), it was shown that the lipid level varies between 1.5-2-7%. It is at a minimum in the larval instars, increases to a maximum in 5-10 day old adults and drops once again after the fifteenth day of adult life. The latter drop is more precipitous in females than males and the authors suggest that the hemolymph lipid may be used for egg maturation since sexual maturity is attained between days 5-10 of adult life. This is a reasonable assumption that should be tested by employing castrated females. However, this tells us little of lipid absorption or transport, as these changes may also be a result of lipid release from and not to the fat body. Figures 5 and 6 demonstrate the pH optima and time course of hydrolysis of TGL by a supernatant fraction separated from the midgut of P . americana. The time course approaches zero order kinetics for the first two hours. There have been several studies on the digestive lipase of this cockroach, among which are those of Wigglesworth (1928), Eisner (1955) and Treherne (1958). Our finding (Gilbert et al., 1965) that the lipase of the mid-gut displays dual pH optima (PH 5.0 and pH 7.2) is of some interest. Wigglesworth (1928) had shown that the gut lipase of this insect was maximally active at an alkaline pH while Eisner (1955) demonstrated it to be quite active even at pH 5.0 (the normal pH of the crop). In general, mammalian pancreatic lipase displays an alkaline pH optimum, although dual acid and alkaline pH

100

LAWRENCE I . GILBERT

optima have been reported for mammalian adipose tissue lipase (Lynn and Perryman, 1960). The greater activity of the mid-gut lipase at pH 5.0 is reasonable since this approaches the in vivo environment. The enhancement of activity by calcium ions compares well with the response of pancreatic lipase to this activator (Gilbert et al., 1965). The similarities between the mammalian and insect digestive lipase would lead us to assume at this point that the absorption products from gut to hemolymph are probably similar to those in the mammals.

2

4

5

6

7

8

9

1

0

PH

FIQ.5. Effect of pH on the hydrolysis of triolein by the digestive lipase of P.americana. Lipolytic activity is expressed in ctslmin in FFA released per mg protein per hour. (From Gilbert et al., 1965.)

At least one species of insect,Galleriamellonella, has a special problem regarding lipid ingestion. The waxmoth larvae invade bee combs and feed upon the wax. The method by which the larvae extract the essentials from the wax and digest them has been an object of experimentation by Niemierko and his colleagues for over a decade. Unfortunately the problem remains unsettled although progress has been made. Niemierko and Wlodawer (1950, 1952a, b) suggested that the rapid diminution of unsaponifiable material and the constant fatty acid level in the larval body during an initial period of starvation is due to a transformation of unsaponifiable matter to fatty acid in the gut. As fatty acid is meta-

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101

bolized, more unsaponifiable material is converted. The unsaponifiable substances containing hydrocarbons and alcohols may be transformed in part by the intestinal flora, but these workers believe that the larval enzymes play a predominant role. During the growth of GaZZeria larvae, glycerides are accumulated and the phospholipid (PL) concentration increases, but at a rate below that of the neutral lipid (Niemierko, 1952). About 47% of the total lipid in the larval intestine is PL (Wlodawer, 1956) although the honeycomb on which the larva feeds is devoid of PL. The PL concentration drops dramatically during starvation and Wlodawer postulates that the products of the hydrolytic 3000 r

L 180

0

60

120

Time (min)

FIG.6. Time course of hydrolysis of tnolein by P. arnericano digestive lipase. Lipolytic activity is expressed in ctslmin in FFA released per mg protein per hour. (From Gilbert et al., 1965.)

breakdown of wax during digestion penetrate the gut wall in a phosphorylated form. An enzyme in the gut wall is presumed to split the PL. Thus, PL most likely play a role in the absorption and transport of ingested lipid in this insect (Niemierko, 1959). This theory gains support from the recent work of Young (1964a, b). In addition, Young found that the Galleria larva has the capacity to convert the iso- or hydroxyacids of the honeycomb into more common (palmitic and oleic acids are predominant) saturated and unsaturated fatty acids. This wax utilization is a most intriguing problem but is easily obscured by the presence of symbiotic micro-organisms in the digestive tract which may be in part responsible for this phenomenon. Subsequent to feeding and absorption of lipid (in some unknown form) into the hemolymph, it is proposed that the fatty acids are then

102

LAWRENCE I . GILBERT

reconstituted into triglycerides in the fat body (Chino and Gilbert, 1964, 1965a). This is not merely supposition but is based on two types of experimental approach. First, TGL as mentioned previously are the major lipid store in the fat body and the hemolymph does not have the capacity to synthesize TGL (Chino and Gilbert, 1965a). Secondly, there have been many in vivo and in vitro experiments that demonstrate that labelled fatty acids injected into the hemolymph or incubated with isolated fat body are to a great extent recovered in the TGL fraction (see Section IV). These fat body triglycerides are then stored in the fat body until metabolic demands are made upon the animal (e.g. flight, molting) whereupon these storage lipids are released into the hemolymph and made available to the appropriate tissue. B . LIPID RELEASE A N D T R A N S P O R T

We have a great deal more information on the release of lipid from the fat body into the hemolymph than on how the ingested lipid finally reaches the storage stage. When in vivo measurements are taken of the lipid concentration in the hemolymph it is clearly difficult to discern between fatty acid influx into the fat body for storage as glycerides, and lipid release from the fat body during the mobilization phase. One approach is to separate the digestive tract-hemolymph system from the fat body-hemolymph system by in vitro studies. When working with an in vitro system it should always be borne in mind that extrapolation back to the whole organism may not necessarily reveal a linear relationship. The ingestion of lipids or conversion of other substrates into lipoidal form, the breakdown in the digestive tract, the reconstitution in the fat body would all be to no avail unless the insect had an efficient mechanism for transporting this potential ATP to a body site where it could be utilized. The most general energy consuming processes in insects are movement (especially flight), oogenesis, embryogenesis and molting (particularly at metamorphosis). How does lipid reach the maturing oocyte or embryo or the flight muscle? What happens once it arrives at the site of utilization? These are the questions that will be at least partially answered in this section. Before turning to the insect, let us look at the situation in the mammal so that parallels can be drawn if indeed they exist. At the pH of mammalian plasma, less than 1% of the FFA would be dissociated, most being bound to protein (Fredrickson and Gordon, 1958) and perhaps the same is true for insect hemolymph. Although each albumin molecule

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103

in mammalian blood bears two sites with a very high affinity for fatty acids, the mechanism of binding remains unknown. Fredrickson and Gordon (1958) suggest that the union is probably electrostatic between the carboxylate ion of the fatty acid and a positively charged group on the protein surface. We know that insect hemolymph contains some FFA as well as protein with an electrophoretic mobility approaching albumin, but are as yet completely ignorant of how and in what form FFA are transported. In mammals, the unique problem of how lipid with its very low solubility in aqueous solutions is transported, is solved by circulation in the form of high or low density lipoproteins or as chylomicrons. The chylomicrons are emulsified particles of lipid in the lymph or blood and are comprised of three zones; a central droplet composed mainly of TGL; a covering of surfactant material (e.g. phospholipid, cholesterol, traces of fatty acid) ; an outer atmosphere of loosely absorbed protein (Dole and Hamlin, 1962). In all of the transport forms in mammals, the major component is TGL although some cholesterol has been identified in all of the lipoprotein complexes. It may be that cholesterol plays a vehicular role in mammals. The physical structure of the lipid droplets in insect hemolymph remains an open question due to the lack of work on the problem. However, we are not completely ignorant of how lipid is transported in insects. An electrophoretic analysis of the hemolymph of P. americana nymphs (Siakotos, 1960a,b) resulted in the finding that three of the five plasma fractions are lipoproteins. Two of these fractions contained protein, phospholipid and carbohydrate while the third had a high neutral lipid and sterol content. Siakotos suggested that the presence of a /?-like lipoprotein may be important in the transport of lipid to and from the reserve tissues. This supposition was supported by the experiments of Tietz (1962) who showed that the locust fat body would synthesize glycerides when incubated in oitro with [14C]palmitate. When this labelled fat body was then incubated in fresh unlabelled medium, glycerides were released into the medium. The specific activity of the released glycerides was about 10 times greater than that present in the fat body. She suggested that in uioo, glycerides are likely mobilized from the fat body into the hemolymph where they are transported as lipoproteins. This suggestion has received extensive confirmation through our work (Chino and Gilbert, 1964, 1965a), and we have identified the form in which the lipid is transported as well. Our studies indicate that diglyceride-hemolymph protein conjugates are the means by which lipid is most likely transported in insects. Although most of the experiments were o n pupae and adults of 5-A.I.P.

4

104

LAWRENCE I . GILBERT

H . cecropia, critical experiments utilizing P . americana and Melanoplus differentialis yielded data indicating the same conclusion. Because of the import of these findings based on electrophoretic, thin layer and column chromatographic techniques as well as radioassay, the results and interpretation of this two-year study will be discussed. In addition, some information on the lipid composition of fat body and hemolymph will also be offered. Table V reveals that the lipid content of adult Cecropia fat body is almost three times greater than that of pupal fat body. In both cases, almost 98% of the glycerides are TGL with the remainder being TABLE V Content of lipid in fat body and hemolymph of H . cecropia Male diapausing pupa Fat body Hemolymph 500 mg 1000 mg (fresh weight) (fresh weight) Content Content

Total lipid 1 . Triglyceride 2. Diglyceride 3. Monoglyceride 4. Sterol Sum o f I , 2, 3, and 4

Male adult ~Fat body 500 mg (fresh weight) Content . _ _ ~

mg

%

mg

%

mg

%

70.0 66.5 0.9 < 0.5 0.8

100 95.0 1.3 0.7 1.1

3.9 0.7 2.6 < 0.2 0.3

100 17.9 66.6 5.1 7.7

233.0 224.7 5.4 < 0.5 1.3

100 96.4 2.3 0.2 0.6

68.7

98.1

3.8

97.3

231.9

99.5

From Chino and Gilbert, 1965a.

diglycerides (DGL) and monoglycerides (MGL). However, in the pupal hemolymph, about 75% of the neutral lipid is DGL, and because of this extraordinary finding we conducted several experiments to ensure that this fraction was indeed DGL (Chino and Gilbert, 1965a). When fat body is incubated with [14C]palmitate, or this fatty acid is injected into pupae, we obtain fat body with high specific activity in the glyceride fraction, the major portion being labelled TGL. This fat body is termed pre-labelled fat body in these experiments. Hemolymph that, had been incubated with this pre-labelled fat body was extracted and the lipid applied to a Florisil column. As shown in Fig. 7, almost all the radioactivity was recovered in the 25% ether fraction, the fraction containing DGL. Standard [14C]DGLwas then mixed with the labelled

_

L I P I D M E T A B O L I S M A N D F U N C T I O N I N INSECTS

105

hemolymph lipid and the whole applied to the Florisil column. Figure 7 demonstrates that peaks of the standard and the labelled hemolymph lipid, and of the mixture of the two, are all eluted at the same time. Though it is possible that sterol and DGL overlap in this chromatographic procedure, we demonstrated through a series of chemical techniques that the radioactive lipid in the 25% ether fraction is not

n

5000r

4000

c

1000

1

I

I!' TGL

-A-&A-A .

0

Herone+5%

4-&

10 ether b 1 5 % ether in hexane in hexone

20

30

Fraction number 2 5 % ether in h e x a n e - t f + 2 % 4 methanol in ether

4% acetic acid in ether

FIG.7. Chromatographic separation of I4Clabelled hemolymph lipid from H . cecropia. I4Clabelled hemolymph lipid. A-A 14C labelled standard diglyceride.0-0 Mixure of standard 14C labelled diglyceride and I T labelled hemolymph lipid. (From Chino and Gilbert, 1965a.)

-.-

cholesterol or any other sterol that forms a digitonide. Finally, thin layer chromatography before and after saponification of the labelled hemolymph lipid demonstrated conclusively that the major lipid in the hemolymph is indeed DGL. This has been since confirmed in L. migrutoriu by Dr. A. Tietz-Devir (personal communication). Figure 8 shows the time course of [I4C]DGLrelease from pre-labelled adult Cecropia fat body into pupal hemolymph. The release of DGL continued through the incubation period whereas the release of TGL

I06

LAWRENCE I . GILBERT

was negligible. Furthermore, this experiment demonstrates that the release of DGL is not a simple “leaking” since only a minute amount is released into phosphate saline. That the release of DGL is rather specific for insect hemolymph is seen in Table VI. This table also shows that [14C]palmitate initially taken up by the fat body is also released into the hemolymph as FFA. However, this release was not specific for insect hemolymph but was also enhanced by serum albumin. 50,OOC

JO.CO0 I

.E. ”7

”

+

1 ~

30,OOL‘

a,

m

m 0 ~

* *

v)

?

; 20,000

-

> -

w

U

u

10,000

0 Incubation lime (min)

FIG.8. Time course of I4C glyceride release from male adult H . cecropia fat body into the hemolymph in uifro. )-u Diglyceride released into hemolymph. -0 Diglyceride released into phosphate saline. A--A Triglyceride released into hemolymph. (From Chino and Gilbert, 1965a.)

The labelled TGL was also released into all incubation media suggesting that the labelled TGL recovered from the incubation media may have “leaked” from the fat body during incubation. In fact, when the pre-labelled fat body was incubated in undiluted hernolymph, TGL release was extremely low. To determine whether DGL release occurs in insects other than Cecropia, similar experiments were conducted with the fat body of the grasshopper and cockroach. We showed that [14C]DGL was rapidly released from the fat body of these insects and that Cecropia hemo-

107

L I P I D METABOLISM AND F U N C T I O N I N INSECTS

lymph stimulated this process as much as the hemolymph of the corresponding insect. Recent work regarding the synthesis of TGL from long chain fatty acids in mammals indicates that DGL is an intermediate TABLE VI Release of glycerides from the fat body of H. cecropia Expt. Incubation No. medium

Triglyceride in medium

% released

Diglyceride in medium cts/min 7; released

Free fatty acid in medium cts/min % released ~.

0.2 ml adult hemolymph 0.2 ml chilled 1 pupal hemolymph 0.3 rnl 5% serum albumin 1 ml phosphate saline 0.3 ml diapause pupal hemolymph 2 I ml diapause pupal hemolymph 1 ml phosphate saline 0.3 ml chilled pupal hemolymph 1 ml chilled pupal hemolymph 0.5 ml rat plasma 0.3 ml rat plasma 1 ml phosphate saline ~~~

~~

944

1.7

91.240

37.7

3,006

4.9

830

I .5

63,180

26.1

2,120

3.5

550

1

.o

2,463

1.7

9,250

15.1

60 I

1.1

2,617

1.8

1,208

2.0

734

0.8

100.070

28.2

I82

0.2

106,530

29.8

3,160

10.9

665

0.7

4,885

1.4

1,110

3.8

1,300

1.4

64.635

28.1

-

150 1,140 1,005

0.2 1.2

29.1 1.8 1.6

.~ .

1.1

66,620 4,120 3,580

1.1

3,225

1.4

995 ~

.-

~~~

~~

Pre-labelled fat body was incubated with incubation media listed below in the presence of 1-2 pmoles GSH at 25" for 60 min. Phosphate saline (pH 6.7) was added to I ml final volume. Expt. 1 :I70 mg pre-labelled fat body containing 54,900 cts/min of TGL, 242,400 cts/min of DGL, 61,200 cts/min of FFA. Expt. 2:180 mg pre-labelled fat body containing 94,100 cts/min of TGL, 357,200 cts/min of DGL, 29,200 cts/min of FFA. Expt. 3:150 mg pre-labelled fat body containing 89,500 ctslmin of TGL, 229,700 ctslmin of DGL, 19,100 cts/min of FFA. (From Chino and Gilbert, 1965a.)

108

LAWRENCE I. GILBERT

and that ATP is required (see Section IV). If the synthesis of TGL and DGL from labelled palmitate by Cecropia fat body proceeds by a similar mechanism, one would expect inhibition of glyceride synthesis by cytochrome inhibitors, uncouplers and other substances that interfere with oxidative phosphorylation. Our data revealed that inhibitors such as KCN, NaN, or 2,4-dinitrophenol prevent the normal incorporation of palmitate into glycerides by the fat body and result in an accumulation of FFA in the tissue. One may ask whether the release of DGL is an active process and may also be affected by these inhibitors. When pre-labelled fat body was incubated with pupal hemolymph in the presence of the above inhibitors, there was a marked inhibition of release of [14C]DGL. In contradistinction, FFA were released at an accelerated rate. Further work led to the conclusion that these inhibitors accelerate the release of FFA from the fat body into the hemolymph by interfering with the incorporation of FFA into neutral lipid. Since the hemolymph itself cannot synthesize glycerides from labelled palmitate, we can exclude this phenomenon as contributing to the DGL content of the hemolymph. Finally, we incubated pupal hemolymph and pre-labelled fat body, and subjected this hemolymph to paper electrophoresis. As illustrated in Fig. 9, four distinct protein bands were resolved, three of which were lipoproteins. Subsequent determination of the distribution of radioactivity of [14C]DGLon the strip clearly revealed that almost all of the DGL was concentrated and recovered in the lipoprotein fraction having a relative mobility of 0.13. From these experiments we can conclude that DGL is rapidly and specifically released from the insect’s fat body into the hemolymph and that very little or no TGL release occurs. This DGL release occurs in uiuo as well as in uitro (Chino and Gilbert, 1964). When [14C]palmitate is injected into a Cecropia pupa, this labelled fatty acid is first incorporated into glyceride by the fat body and subsequently released as DGL. Consequently, the specific radioactivity of the glycerides in the hemolymph is much greater than that in the fat body because TGL is the major component of the neutral lipid in the fat body. In other words, DGL with an extremely high specific radioactivity is continuously being released into the hemolymph, while the low specific activity TGL remains for the most part in the fat body. The demonstration of DGL release from the fat body of four species of insects (silkmoth, grasshopper, locust, cockroach) suggests that this may be a phenomenon common to many or all insects.

Distonce from o r i g i n ( c m )

FIG.9. Separation of H . cecropia hernolymph proteins and associated 14C labelled diglycerides by paper electrophoresis. Ordinate: cts/min of ["C] diglyceride found in 1 cm piece of electrophoretogram. LP is strip stained by Sudan black, lipoprotein. P is strip stained by brornophenol blve, protein. (From Chino and Gilbert, 1965a.)

110

LAWRENCE I . GILBERT

In addition the data suggest that the release of DGL is an endergonic process and requires ATP as is the case in glyceride synthesis. In contradistinction, the release of FFA seems to follow a concentration gradient and depends on the level of FFA in the fat body. This then is a passive process. Thus, we might expect that in an actively metabolizing tissue like adult Cecropia fat body, DGL release is high, while FFA release is high in tissues of lower metabolism such as the fat body of a diapausing pupa. How does this compare with the transport mechanism in mammals to which we have already alluded? In mammals, the main components in the transport of long chain fatty acids are most likely TGL and FFA. In fact, the major neutral lipid component in mammalian plasma is TGL with DGL and MGL present as only minor constituents. This differs from the insect case in that DGL replaces TGL in the insect. In mammals, the TGL found in the plasma is conjugated with plasma proteins as the low density lipoprotein, chylomicron, etc. It appears that in the hemolymph of the insect, DGL is carried by, and perhaps conjugated to, one of the hemolymph proteins. These findings indicate that DGL-protein complexes and FFA (perhaps also bound to protein) are the means by which insects transport lipid. That acetate can be oxidized by insect muscle is now taken for granted since it was demonstrated almost 20 years ago (Barron and Tahmisian, 1948). In 1955, Sacktor suggested that stored lipid is broken down to acetate in the fat body of Musca domestica and transported in that form to the flight muscle. The work discussed above demonstrates that this is not true for several insect forms although we have not yet completed our investigation of the Diptera. The physiological significance of the transport of DGL will be discussed in Sections 111 and IV. At this time, we can say that in the Cecropia moth for instance, the flight muscles utilize long chain fatty acids almost exclusively but store only little lipid. Thus lipid must be transported to this active tissue perhaps in the form of DGL. In addition, long chain fatty acids are necessary for normal oogenesis and must be transported from the fat body to the ovaries in a similar way. Furthermore, of importance is the fact that DGL are intermediates not only in TGL synthesis but also in phospholipid synthesis (Fig. 15; see Section IV). C . EXTRA-DIGESTIVE LIPASES

Prior to discussing the final utilization of the transported DGL as a source of energy, we should consider one other problem in the sequence

L I P I D METABOLISM A N D F U N C T I O N IN INSECTS

111

of events beginning with the ingestion of lipoidal material and ending with the oxidation of fatty acids. If lipid is stored as TGL within the fat body and subsequently released as DGL, it follows that at least one fatty acid moiety must be cleaved from each TGL molecule in the fat body before transport can occur. Until very recently, no true extradigestive lipase has been demonstrated in insects by the use of biochemical procedures. Histochemical work has demonstrated the presence in the fat body of an enzyme that will hydrolyze chemical such as a-naphthyl acetate (see for example Wigglesworth, 1958), but these histochemicals are certainly not the true substrate in the insect. There have been a large number of reports in the past by Prof. J. C. George and his group on the ability of various insect (and vertebrate) tissues to hydrolyze tributyrin (Bhakthan, 1964; George and Bhakthan, 1960a, b, 1961, 1963; George and Eapen, 1959a, b; George et a/., 1958, etc.). Although this work is of real interest, it does not elucidate the action of a lipase, contrary to what the authors state. Although their work demonstrated that honey bee flight muscle can oxidize butyrate, this C, fatty acid is more likely present in the insect as a result of the /3-oxidation of longer chain fatty acids (see Section 1II.D.) rather than of the hydrolysis of tributyrin. It is more likely that most of the work of the Indian group is conducted on esterases such as that described by Van Asparen (1959). She showed the presence of potent ali-esterase activity in various body regions of Musca domestica. The enzyme(s) was extremely efficient in hydrolyzing both triacetin and tributyrin. Of interest is the fact that Van Asperen found 22% of the esterase activity responsible for the hydrolysis of tributyrin to be localized in the head of the insect. Since the head contains little or no fat body, but is composed mainly of nervous tissue, the enzyme described is most likely not a true lipase. The ability of an enzyme to cleave butyrate from tributyrin may be far removed from its primary action. Our laboratory has been engaged in an attempt to characterize a true extra-digestive lipase in various stages of the life cycle of H . cecropia for several years. Almost every conventional technique has been employed including titrimetric determination, spectrophotometric assay and manometry. In each case no true lipase could be demonstrated, but potent esterase activity was found in several tissues. Table VII reveals that hemolymph, fat body and gut from several developmental stages show no lipase activity when assayed by manometFy. All tissues are capable of hydrolyzing triacetin, with gut exhibiting the greatest

112

L A W R E N C E 1. G I L B E R T

activity. In both the pupal and adult stages, the male fat body shows more active esterase than that of the female, and the adult tissues are far more potent in splitting the ester bond than the respective tissues of the pupa. In no case could we detect the release of FFA from the water-insoluble glycerides (olive oil). Since the flight muscle is the most metabolically active tissue in the moth we believed that it might be the source of the enzyme we were seeking. However, repeated attempts to demonstrate a true lipase in TABLE VII Esterase activity in tissues of If. cecropia Hydrolytic activity Stage

Enzyme source

(pl C 0 2 / m g protein

N/hr) Triacetin Chilled male pupa Chilled female pupa 20-Day-old male Developing adult 1-Day-old male adult 1-Day-old female adult

whole blood fat body homogenate gut homogenate whole blood fat body homogenate gut homogenate whole blood fat body homogenate fat body homogenate fat body homogenate

6.02 266.34 890.79 2.40 56.54 375.70 16.49 27 1.57 408.50 266.70

. _ _-

Olive oil

0 0 -~

0

0 0 0 0

Manometric assay. Reaction mixture contained 1 ml substrate; I ml enzyme source; 1 ml bicarbonate buffer. Triacetin added as 10% solution in buffer. Olive oil added as 10% emulsion in buffer prepared in Waring Blender with 1% polyvinyl alcohol. Run at 25", pH 7.4. (From Gilbert et a/., 1965.)

this tissue failed. The homogenates did split the ester bonds of tributyrin and ethyl acetate, but not those of higher fatty acid glycerides. The conditions of the assay were not limiting since the reactions that did occur were linear with time. The optimum pH for flight muscle esterase is about pH 7.4 and activity rapidly drops when assayed one pH unit below this value (Gilbert et al., 1965). T h w various tissues from Cecropia possess the ability to hydrolyze water-soluble esters, but showed no lipase activity. All our other data (see previous section) indicated that this insect, obviously utilizing products of glyceride hydrolysis, must possess a means of converting the storage lipids to a

L I P I D METABOLISM AND F U N C T I O N I N INSECTS

113

utilizable state. If the lipase present was only minimally active, perhaps the available assays were below the level of necessary sensitivity. We therefore developed a highly sensitive assay for lipase activity utilizing [14C]labelledglycerides (Chino and Gilbert, 1965b). Utilization of this sensitive assay provided proof for the existence of a true lipase that hydrolyzes triolein in the fat body of the Cecropia moth (Gilbert et al., 1965; Figs. 10 and 11). Table VIII indicates the activity of lipase from several tissues on the DGL and TGL. It is TABLE VIII Hydrolysis of di- and triglycerides by insect tissues

Enzyme source

Substrate

Pupal Cecropia haemolymph Adult male Cecropia fat body

l4C-trio1ein 14C-triolein 14C-diolein 14C-triolein '*C-diolein l4C-triolein '4C-triolein l4C-dio1ein

Cecropia flight muscle Cockroach fat body Cockroach midgut

Hydrolytic activity (cts/min in unesterified fatty acid released from substrate/mg protein/hr)

0 35,26 41,46 41,54 290,203 33 973 1016

Run in tris buffer, pH 7.2 at 30". The substrate is actually a mixture of labelled glyceride plus unlabelled carrier glyceride. The enzyme source is the middle layer from a centrifuged homogenate. Assayed according to Chino and Gilbert (1964b). (From Gilbert et a/.,1965.)

important to note that the lipase from Cecropia flight muscle was five times more active against diolein than triolein. This was in contrast to results with other tissues including Cecropia fat body. It is known that calcium ions activate mammalian pancreatic or intestinal lipase (Desnuelle et a/., 1950; DiNella et al., 1960; Chino and Gilbert, 1965b). Our results (Gilbert et a/., 1965) demonstrate that calcium ions stimulate the cockroach digestive lipase threefold but have no effect on the extra-digestive lipases. The data indicate that definite lipolytic action is associated with Cecropia fat body, but the activity is low. To investigate the possibility that an endogenous lipase inhibitor resides in the fat body, attempts were made to remove this hypothetical inhibitor by preparing acetone powders of the fat body lipase and dialyzing fresh

114

LAWRENCE I. GILBERT

I

I

4

8

I I2

Tlme ( h r )

FIG. 10. Time course of hydrolysis of triolein by adult male H. cecropia fat body. Lipolytic activity is expressed in cts/min in FFA released per mg protein per hour. p H 7.2 in tris buffer. (From Gilbert et a/., 1965.)

I

I

6

7

I

I

I

5

8

9

P"

FIG. 1 I . Effect of p H on the hydrolysis of triolein by the fat body lipase of adult H. FFA released per rng protein per hour. (From Gilbert e f a/., 1965.)

cecropia. Lipolytic activity is expressed in cts/rnin in

L I P I D METABOLISM A N D F U N C T I O N I N I N S E C T S

1 I5

extract against water. In neither case was the enzymic activity enhanced. Furthermore, addition of fat body homogenate to incubating mammalian pancreatic lipase did not diminish the activity of the mammalian enzyme. It thus appears that no endogenous inhibitor is present. Thus we have shown that extra-digestive lipases do exist in insect tissues but that their activity is not measurable (at least in H . cecropia) when conventional techniques are used. Techniques such as manometry do, however, uncover esterase activity when substrates such as triacetin and tributyrin are used. The distinction between lipases and esterases that hydrolyze short chain fatty acid glycerides is certainly not clear, but several criteria can be used to differentiate between these two classes of glycerol ester hydrolases. In general, a lipase has a preference for long chains in the acyl groups of the glycerol ester (as in triolein, tripalmitin, etc.) whereas an esterase has a preference for short chains in the acyl groups (as in triacetin, tributyrin, etc.). When the ester substrate is soluble in water it is not attacked by pancreatic lipase (Desnuelle and Savary, 1963). These authors assume that an emulsion contains multimolecular aggregates estranged from the aqueous medium by an interface. The lipase molecule is absorbed to this interface and can thus exert its effect. In aqueous solution, the lipase would be inactive. In fact, in mammalian adipose tissue, different enzymes are responsible for the hydrolysis of long and short chain fatty acid glycerides, one of which has been termed a “tributyrinase” (Schnatz and Williams, 1962). Finally, the use of substrates such as triacetin and tributyrin has little physiological significance since these substances are not usually present in animal tissues, although the rate of hydrolysis of these esters may be an indicator of general metabolic activity . In Cecropia, esterase activity in the tissues studied increases steadily during the pupal-adult transformation and reaches its highest level in the adult moth (Gilbert et a/., 1965). This increase during development is most likely a reflection of the general increase in metabolic rate during adult development. The sexual dimorphism in esterase activity is another example of biochemical differences between males and females of this species (see Section I I ) , and again may be related to the more active flying conducted by the male moth. The physiological significance of the ability of these tissues to hydrolyze triacetin and tributyrin remains obscure. Our results on the flight muscle esterase are in agreement with those of Fodor (1948) who showed that locust flight muscle could not hydrolyze olive oil but didcleave the fatty acid moiety from monobutyrin and methylbutyrate. Cecropia flight muscle

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LAWRENCE I . GILBERT

is capable of hydrolyzing tributyrin at about the same rate as S.gregaria flight muscle (George et al., 1958) and honey bee flight muscle (George and Bhakthan, 1963). It is possible that the enzymes in various insect tissues responsible for the hydrolysis of these short chain fatty acid glycerol esters are in reality primarily concerned with completely different reactions and that glycerol ester hydrolase activity is only a secondary effect. Desnuelle and Savary (1963) suggest that the hydrolytic activity of some of the so-called pancreatic esterases merely corresponds to side activities of well known proteolytic enzymes. The most significant results on this extra-digestive lipase of Cecropia are those indicating that Cecropia flight muscle preferentially hydrolyzes DGL whereas the other lipases examined preferentially attack TGL. This question of lipase in flight muscle and fat body is an integral part of the general problem of lipid transport. We must assume that the lipase action of the fat body results in DGL that are then transported in the hemolymph as discussed previously, to sites of utilization such as the flight muscle. We must further assume that the DGL are separated from their carrier protein as they move into the tissue just as fatty acids disengage from their carrier protein as they enter mammalian tissue (Fredrickson and Gordon, 1958). The data suggest that DGL enters the flight muscle intact, whereupon the fatty acid is cleaved from the glycerol ester and rapidly oxidized in the flight muscle sarcosomes. Thus, ingested lipid makes its way to the tissue where it can finally be completely oxidized and contribute to the energy currency of the cell. The next question is how this lipid catabolism takes place. D . FATTY ACID CATABOLISM

The major portion of our knowledge regarding fatty acid oxidation in insects comes from the study of flight muscle. Because of this, flight muscle will serve as our example of an active tissue capable of converting fatty acids to CO, and water. Before discussing this wide subject, let us look at some other processes where fatty acid oxidation may play a major role. 1. Embryos

Almost 30 years ago, Ditman and Weiland (1938) proposed that reduced body water, increased lipid and increased saturation of the lipid are associated with the ability of insects to withstand hibernating conditions (i.e., diapause). This has been consistently confirmed in the last decades and indicates that fatty acid oxidation may play a large

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117

role in supplying the energy required for the relative long diapause periods of some insects. In several insects at least, free lipids are the primary source of energy for the larval-pupal moult as in Tenebrio molitor (Moran, I959), Tribolium confusum (Villeneuve and Lemonde, 1963) and possibly Bombyx (Niemierko, 1959). In addition, as we have seen previously (Section II), lipids are a prime source of energy for insect embryogenesis. In the case of the Cecropia egg for example, there is a dramatic reduction in lipid content between the time of egg laying and the hatching of the first instar larva. Lipids are known to be catabolized to a greater extent in cleidoic eggs than in acleidoic eggs (cf. Babcock and Rutschky, 1961). In 1885, Tichomirov showed that there is a 20% decrease in the free lipid from the termination of diapause to hatching in the eggs of B. mori. Similar studies have been made on other insect eggs, notably M . diferentialis which combined with manometric measurements suggested that about 90% of the oxygen consumed by Melanoplus embryos was used for lipid catabolism (Boell, 1935; Hill, 1945; Slifer, 1930). Further studies on other species led to similar conclusions although only a small drop in lipid was noted during the embryonic development of Oncopeltus (cf. Babcock and Rutschky, 1961). For example, Allais et a/. (1964) reported a 32% decrease in the lipid content during the 18 days of embryogenesis of L. migratoria, much of which was due to the catabolism of glycerides. In Dixippus, about two-thirds of the energy used for embryogenesis is attributed to lipid. About one-half of the original lipid is oxidized to CO, and water during embryonic development as compared to onefourth of the protein (Lafon, 1950). Recent studies on the ovoviviparous cockroach, Leucophaea maderae (Gilbert, unpublished observations) reveal a dramatic increase in the lipid content of the ovary of an adult female prior to oogenesis (about 0-5 mg) to approximately 80 mg in an ovary containing mature eggs just prior to ovulation. This is associated with a concomitant decrease in the lipid content of the fat body indicating transport of lipid to the active ovary. After fertilization, the embryos are enclosed within an egg case held in the female’s uterus and embryogenesis proceeds for about two months. During this period, the content of lipid decreases from about 80 mg per egg case to approximately 40 mg per egg case, indicating that lipid is one major substrate for embryogenesis in this insect. This supposition is supported by our radiotracer studies in which ovaries undergoing oogenesis were found to possess a profound capacity to incorporate [l*C]palmitate into egg .lipids (L. I . Gilbert, unpublished observations).

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LAWRENCE I . GILBERT

There have been relatively few studies on the lipid composition of insect eggs or embryos. One of the exceptions is the recent work of Strong (1963a, b, 1964) who demonstrated that 13.9% of the fresh weight of the aphid egg is lipid. Of this, 85% is neutral lipid, 2% FFA, 4.75% sterol ester and 7.5% is made up of sterols, hydrocarbons and miscellaneous lipids. Of the fatty acids composing the neutral lipid fraction, he has demonstrated a preponderance of myristoleic (C,,,,), myristic and oleic acids. Our studies on the developing eggs of L. maderae (L. I. Gilbert, unpublished observations) demonstrated that the egg lipid is made up of 9O-92% “neutral lipid” and 8-10% phospholipid. In the former fraction, about 90% is TGL, 2 - 3% FFA, 1 - 2% sterol and about 2% sterol ester and hydrocarbon. The remaining lipid is MGL and DGL. During embryogenesis, an increasing percentage falls into the phospholipid fraction (up to 16% of the total lipid) but no drastic qualitative changes occur in the neutral lipid fraction. The mechanism of lipid utilization by embryos has not been comprehensively studied with biochemical techniques, although a number of histochemical investigations have been reported. For example, Nath et al. (1958) demonstrated the presence of three types of lipid bodies in the oocyte of P . americana. One was believed to be wholly phospholipid, one TGL surrounded by an incomplete sheath of PL, while the third is composed of TGL. One can speculate that the TGL may be an energy source for oogenesis while the PL may be used as basic material for membrane formation and microcompartmentalization of the cell. While on the subject of insect eggs, it is of interest to note that in 1952, Hsu reported that mitochondria containing large quantities of unsaturated lipid pass from the Drosophilu nurse cell to the oocyte. According to this author, these mitochondria then transform into lipid yolk bodies. As we shall see subsequently, if any subcellular organelle is to be a lipid body precursor, the mitochondria containing a large quantity of lipid are almost ideal candidates. The notion that the mitochondria may act as an ultimate source of energy as well as an energy converter is a novel (and perhaps far-fetched) idea. As seen from the foregoing, the role of lipid in insect development poses some intriguing questions. Inroads have been made but the fundamental questions remain unanswered. This is in contrast to the more recent attempts to elucidate the role of lipids in flight muscle metabolism. 2. Flight muscle metabolism The flight muscle of locusts, moths, flies and bees has contributed greatly to many aspects of our understanding of insect biochemistry

LIPID METABOLISM A N D F U N C T I O N I N INSECTS

119

(cf. Gilbert and Schneiderman, 1961a). This is especially true regarding the oxidation of fatty acids, in which case muscle appears to be the sole object of contemporary experimentation. A great deal of the research on insect flight muscle has been concerned with the a-glycerophosphate dehydrogenase system in those insects that use carbohydrate as flight fuel (Chance and Sacktor, 1958; Sacktor and Dick, 1962; Van den Burgh and Slater, 1962; Zebe et al., 1959). Although the data are of great general interest, they need not concern us here in our discussion of muscle that preferentially oxidizes lipid. However, it should be pointed out that a-glycerophosphate dehydrogenase can mediate the reduction of dihydroxyacetone phosphate to a-glycerophosphate which in turn can be converted to various complex lipids (see Section IV). In addition, a-glycerophosphate may play a special role in the so-called “a-glycerophosphate cycle” (Klingenberg and Biicher, 1960). This cycle is a mechanism whereby hydrogen generated into the extra-mitochondria1 pool of NADH by the EmdenMeyerhof pathway is made available to the respiratory chain in the mitochondria. The hydrogen is transferred to dihydroxyacetone phosphate by the action of an NAD-dependent extra-mitochondria1 aglycerophosphate dehydrogenase. The a-glycerophosphate diffuses readily into the mitochondrial compartment where a mitochondrialbound a-glycerophosphate dehydrogenase is located. This enzyme is not coupled to NAD but enters directly into the respiratory chain (Biicher et al., 1959; Sacktor, 1959; Klingenberg and Biicher, 1960). Intensive study into lipid utilization by insect flight muscle essentially began with the comprehensive investigation of Weis-Fogh (1952) on flight in Schistocerca gregaria. He found that 80-85% of the total energy expended in the first five hours of flight was derived from lipid. The fat body was responsible for delivering 85-90% of the energy expended by the muscles. Previously, Krogh and Weis-Fogh (1951) demonstrated a decline in RQ from 0.82 to 0.75 during the first 90 min. of flight (see also Bode and Klingenberg, 1964). An RQ approximating 0.7 is indicative of almost exclusive lipid catabolism. Weis-Fogh’s experiments were well conceived, used insects raised under near natural conditions and left little doubt that lipid was the main fuel for locust flight (see also Fulton and Romney, 1940). As Weis-Fogh points out, lipid is an ideal substrate for flying insects since the hydration of glycogen would make isocaloric quantities eight times heavier than lipid (and result in an extremely heavy flying machine). Since Weis-Fogh‘s original observations, a number of publications have appeared that deal with some aspect of the relation of lipids to 6+A.I.P.

4

120

LAWRENCE I. GILBERT

flight muscle metabolism. For the most part, they are concerned with the study of rates of oxidation of fatty acids by flight muscle, flight muscle homogenates or sub-cellular fractions and the assay of particular enzymes of the oxidative pathway. Let us now consider the final step 0

II

:--------,R-CH,-CH,C-OH

+

A

CoASH ATP

0 II R- CH,CH,C-SCoA

HP,O?- + A M P

G FAD

0

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$".

FmH,

R-CH=CHC-SCoA

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JI Krebs' cycle FIG.12. The 8-oxidation of fatty acids. The enzymes involved are: a. acyl thiokinase b. acyl dehydrogenase c. enol hydrase d. 8-hydroxyacyl dehydrogenase e. thiolase

in the utilization of lipid by the insect, namely the oxidation. of fatty acids to carbon dioxide and water. Figure 12 outlines the known steps of the Boxidation of fatty acids as we know them from the classic work with microbial and vertebrate systems. It can be readily seen that a relatively enormous quantity of

LIPID METABOLISM A N D FUNCTION I N INSECTS

121

energy is made available to the cell when long chain fatty acids are completely oxidized. Although one ATP is sacrificed in the “sparking reaction” (reaction a in Fig. 12), only one is utilized regardless of the chain length of the fatty acid. Once the process of /?-oxidationis initiated within the mitochondrion, it usually runs to completion by the removal of C, segments from the original fatty acid, which then enter the Krebs’ cycle as acetyl CoA. For each C, unit the cell gains the potential energy inherent in 1 reduced FAD, 1 reduced NAD and that released in one complete turn of the Krebs’ cycle. For each mole of a saturated C18fatty acid that is completely oxidized, the cell gains more than 140 moles of ATP. As an example of the work conducted on the flight muscle oxidation of fatty acids, let us again consider the Cecropia moth (Domroese and Gilbert, 1964). During the second and fifth day of adult life, male Cecropia moths show a sudden utilization of total lipid which had remained more or less a constant quantity during adult development (see Section 11). This sudden drop in lipid was thought to be due to the rather abrupt assumption of flight activity in young adults. Prevention of flight results in a conservation of lipid, suggesting that flight is the major causative factor in the sudden decrease in lipid content of the moth. The adult moths do not feed and of necessity, the metabolic substrates of the adult are limited to those available after the requirements of metamorphosis have been satisfied. The carbohydrate store of the male moth is almost completely exhausted during adult development leading to the hypothesis that lipid must be the main substrate for adult life (namely flight muscle metabolism; see Table IX). By measuring the endogenous Qo, of flight muscle homogenates we were able to obtain a reproducible estimate of their metabolic rate. Dialysis of these homogenates resulted in a large reduction in the Qoa due to the loss of requisite cofactors. By adding cofactors back to the dialyzed homogenate we were able to determine which were necessary for normal cellular metabolism. These results show that ATP, Mg+ and citrate are required to restore the endogenous respiration of dialyzed homogenates to the level of the non-dialyzed control. This infers that these same cofactors are necessary for fatty acid oxidation. The stimulation of respiration by citrate may be due to some extent to its oxidation as an added substrate or perhaps to its stimulation of oxidation of endogenous lipid by priming the citric acid cycle. To test these two alternatives the oxidation of [14C]palmitate by dialyzed flight muscle homogenate was measured in the presence of added cofactors. The results (Table X) clearly indicate that added citrate stimulates fatty +

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LAWRENCE I . GILBERT

acid oxidation and verify the need for ATP and Mg . The stimulatory effect of citrate is not specific and other citric acid cycle intermediates can also stimulate fatty acid oxidation. The pH optimum for the oxidation of [l4CJpalmitate by the dialyzed homogenate with added cofactors coincides with that of endogenous respiration of undialyzed homogenate (Fig. 13). Malonate was an effective inhibitor of fight muscle respiration and together with the demonstration that citric acid cycle intermediates stimulate fatty acid oxidation, indicates that the pathway of fatty acid +

+

TABLE IX Oxidation of various labelled substrates by homogenate of male H. cecropia flight muscle

Substrate (1.1 x los cts/min) Acetate Alanine Glucose Pyruvate Glycerol Succinate Butyrate Decanoate Palmitate

Acid insoluble dry weight (ms) 10.20

10.10 8.43 8.19 7.85 7.92 8-34 10.40 10.30

1 4 ~ 0 ,

total cts/min 60 min

53,410 16,853 4,063 115 1,581 55,032 43,042 56,071 113,510

cts/min mg acid insol. dry weight

14C02

5,236.3 1,668.6 482-0 14.0 201.4 6,948.5 5,160-9 5,391.4 1 1,020.4

Homogenate prepared in 0 . 1phosphate ~ buffer, pH 7. (From Domroese and Gilbert, 1964.)

oxidation is via the citric acid cycle. Additional evidence for the participation of this pathway was obtained by measuring the production of 14C02from a fatty acid labelled in different numbered carbons. Studies on flight muscle sarcosomes (mitochondria) showed that the same cofactors were necessary if fatty acids were to be oxidized at a maximum rate. [It should be noted that not all insect mitochondria can oxidize fatty acids as can be seen from the numerous studies on flight muscle sarcosomes from carbohydrate utilizing insects and from the study of mitochondria isolated from the silkworm intestine (Sridhara, 1965).] In addition, we studied the fatty acid activating enzyme of the Cecropia flight muscle and found that it was maximally active only when CoA, ATP and fatty acid were present. That relatively high con-

LIPID METABOLISM A N D FUNCTION IN INSECTS

123

centrations of coenzyme A exist in insect tissues is no longer in doubt (Boccacci and Bettini, 1956). The above results showed that fatty acids are actively oxidized by fight muscle homogenates whereas glucose, pyruvate and glycerol were oxidized only to a limited extent, supporting the original observations 17 16

15 14

13 12 L

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400

350

300

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6.0 6.2 6.4 6.6 6.8 7.0 7.2 7.4 7.6

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FIG.13. Effect of pH on endogenous respiration and oxidation of palmitate-l-'*C by flight muscle homogenates from H. cecropia moths. (From Domroese and Gilbert, 1964.)

of Zebe (1954) on a feeding moth. The underlying basis for this inactivity presents an intriguing problem in the comparative physiology of insect muscle. Zebe (1959a, b) has already suggested that insect flight muscle is of three physiological types : the exclusive carbohydrate users, the flies; the exclusive lipid users, the moths and butterflies; and those that

124

LAWRENCE I . GILBERT

can use both fat and carbohydrate, the locusts. Studies on the biochemistry of fly and locust muscle have shown no qualitative differences correlated with the type of fuel used (Zebe, 1959a; Boettiger, 1960). That some differences do exist between the above groups of insects is suggested by the observation that after flight there is a continued elevation of respiration for an hour or more in locusts and moths whereas this does not occur in the Diptera (Chadwick and Gilmour, TABLE X Cofactors necessary for the oxidation of [ l-14C]palmitic acid by dialyzed homogenate of male H. cecropia flight muscle Reaction medium Complete -ATP Cit r at e -MgCL No cofactors Undialysed control Boiled control

-

Acid insol. dry weight (mg)

14C02cts/min 60 min

13.03 13.10 13-04 13.65 12.4 1

67,030 6,220 21,533 9,453 6,376

5,144.3 474.8 1,651.3 692-5 513.8

16-12 12.05

74,150 126

4,599.3 10.5

14COacts/min mg acid insol. dry wt.

The complete reaction medium received 1 ml of dialyzed homogenate, 1.2 x lo5ctslmin of [l-14C]palmiticacid, and 1 p o l e of each of ATP, citrate, and MgCla added in a total ~ buffer, pH 7. Flasks lacking one or more cofactors received of 0-3 ml. of 0 . 1 phosphate an equivalent volume of buffer. Control flasks received 0.3 ml of buffer in addition to the homogenate indicated. (From Domroese and Gilbert, 1964.)

1940; Krogh and Weis-Fogh, 1951; Zebe, 1954). Our findings (Domroese and Gilbert, 1964) that flight muscle of Cecropia cannot appreciably oxidize glucose and glycolytic end products indicates that further experiments relating to the biochemical pathways of this tissue may reveal sigmficant quantitative differences when compared to flight muscle from insects of other orders. In respect to the requisites for fatty acid oxidation, the system we studied shows similarities to the requirements for fatty acid oxidation by rat liver mitochondria (Kennedy and Lehninger, 1949). The insect system differs from the mammalian system by the lack of cytochrome c dependence. This may be due to the naturally high concentration of cytochrome c in moth flight muscle sarcosomes. E. Margoliash (personal

L I P I D METABOLISM A N D F U N C T I O N I N INSECTS

125

communication) found that moth flight muscle contains the highest concentration of cytochrome c yet described. Because of the naturally high concentrations, it may persist in sufficient quantities during tissue preparation, eliminating the necessity of an exogenous supply. The requirements for the flight muscle fatty acid oxidizing system of Cecropia are also similar to those known in other invertebrates. At present the only well characterized invertebrate systems have been described in the hepatopancreas of Carcinus (Munday and Munn, 1962) and the thoracic muscles of the locust (Meyer et al., 1960). The Cecropia system is comparable to the locust particulate system in that both require ATP and magnesium for optimum fatty acid oxidation. It differs, however, in that addition of citric acid cycle intermediates enhances the oxidative capacity of the Cecropia flight muscle. The action of the intermediate in Cecropia is similar to that of the intermediate necessary as a primer in rat liver mitochondria, whereas the primer in the oxidation of higher fatty acids by the locust thoracic particle is butyrate. Butyrate is unable to replace the citric acid cycle intermediate in our system. The pH optimum for oxidation of fatty acids in Cecropia flight muscle was identical with that in the locust (6.4). This is somewhat lower than that found in mammalian systems but is in keeping with the fact that the pH of the hemolymph of various Lepidoptera and locusts has been reported to fall on the acid side of neutrality. The complete &oxidation scheme has not been worked out in detail in any one insect. However, recent work has identified individual enzymes in insect tissue that are similar to those found in mammalian tissue, suggesting the presence of the /3-oxidative pathway. The acetate activating enzyme was detected in the flight muscle of the honey bee (Hoskins et al., 1956). Nelson (1958) showed the presence of fatty acid activating enzymes in the fat body of the cockroach and several species of Lepidoptera. In 1960, Zebe identified the condensing enzyme that leads activated acetate into the citric acid cycle and /3-ketoacyl thiolase that leads /3-ketoacyl into the fatty acid cycle in locust flight muscle. In comparing several enzymes from leg and flight muscle of a carbohydrate utilizing insect (bee) and one utilizing lipid (locust), Beenakkers (1963a) demonstrated that although lactic dehydrogenase activity was high in the leg muscle, it was only slight in the flight muscles of both insects. In all cases, the locust flight muscle homogenate oxidized fatty acids at a greater rate than isolated sarcosomes. The addition of carnitine however, raised the activity of the mitoehondria in relation to that of the homogenate while the addition of palmityl camithe

126

L A W R E N C E I . G I L B ER T

resulted in the mitochondria exhibiting 50% more activity than the homogenate. Beenakkers postulates that under normal physiological conditions, carnitine may activate the fatty acids in insect flight muscle. Carnitine transacetylase mediates the reaction : Acetylcarnitine

+ CoASH +carnitine + CoASAc

resulting in activated acetate. Although 5,150 enzyme units were located in the locust flight muscle, that of the bee was devoid of activity (Beenakkers and Klingenberg, 1964). The enzyme appears to be present in the soluble matrix of the mitochondria since on disruption of these organelles, the enzymic activity becomes associated with the soluble fraction. As Beenakkers and Klingenberg suggest, carnitine transacetylase is not an obligate intermediate for fatty acid oxidation (and possibly not needed in the Cecropia flight muscle) but may function in the transport of fatty acids into and within the mitochondria. Further work is needed to prove this supposition but it gains some support from the recent work of Bressler and Friedberg (1964) who demonstrated that carnitine not only stimulates the oxidation of palmityl CoA but also augments the rate of palmitate incorporation into mitochondrial phospholipids. Carnitine may thus function as an acyl carrier from extramitochondrial to intramitochondrial sites. This entire problem of fatty acid relocation within the mitochondrion is an important one and will undoubtedly receive more attention in the future. Recently, Beenakkers (1963b, c, 1965) reported on the fatty acid content of the fat body, hemolymph and flight muscles of L. migrutoriu before and after fight. He calculates that these tissues contain enough lipid reserve for five hours of flight and if the substrates of other body parts are utilized, the locust can stay aloft for seven to eight hours. During flight, the esterified fatty acid concentration of the hemolymph increases from 4.0 to 16.9 mg fatty acid per ml while the FFA concentration remains the same. There is a simultaneous 30% increase in the fatty acid content of the fight muscles, half of which is due to linolenate. This suggests a transport of fatty acids from the hemolymph to the flight muscle. The fat body contains 43.5% of its dry weight as fatty acid of which 42% is palmitate. The hemolymph yields 37% palmitate while the flight muscle contains 24% of its fatty acids in the form of palmitate. The decreasing order of concentration supports the contention of transport in that direction and final oxidation in the flight muscles. Beenakkers believes that palmitate and oleate are preferentially oxidized by the flight muscle and calculates that during flight the flight muscle consumes some 23 mg fatty acids per gram fresh weight per

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127

hour and that this fatty acid ultimately is derived from the fat body. These data agree with those discussed previously, indicating the use of fatty acids for flight fuel in some insects. Of particular interest is Beenakker's finding that it is the esterified fatty acids which appear to play the predominant role in lipid transport, although he did not elucidate the nature of these esters. It is probable that they are diglycerides as discussed in Section 111. Thus in insects that ingest lipid, we can now trace with some authority the phases of digestion, storage, release from storage centres, transport and finally oxidation of the fatty acids in tissues such as fight muscle. However, it would be rare indeed for an animal to ingest enough lipid to establish a lipid storage supply that may equal 50% of its entire weight. More commonly we would expect the insect to have the capacity to convert non-lipid substrates into lipid and to synthesize long chain fatty acids and then glycerides from simpler precursors (e.g. acetate). These two processes of substrate interconversion and lipid biosynthesis probably play a more important role in supplying lipid than that of lipid digestion. This is particularly true in phytophagous insects since most plants contain only minute quantities of lipid relative to the amount of carbohydrate (i.e., starch and cellulose). IV. LIPID BIOSYNTHESIS A . GENERAL MECHANISM OF FATTY A C I D SYNTHESIS

Lipid biosynthesis can be thought of as a two stage process. First, fatty acids must be synthesized and second, the fatty acids must enter the pathway leading to the synthesis of glycerides. In both cases, we shall use schemes derived from work on organisms other than insects to present the overall picture; and then attempt to discuss those investigations dealing with insect material. Figure 14 reveals the essentials of what is now understood to be the main pathway for the biosynthesis of fatty acids in living things. In an excellent study, Wakil et al. (1964) have demonstrated that a heat-stable protein called an acyl carrier protein (ACP) participates in fatty acid synthesis as well. This ACP acts by accepting acetyl and malonyl groups from their CoA derivatives forming covalently linked acetyl and malonyl derivatives. Acetyl-ACP and malonyl-ACP then react to yield acetoacetyl-ACP. In the presence of NADPH, they react to form long chain fatty acids. When an enzyme fraction is incubated with acetyl CoA, malonyl CoA, NADPH and ACP, a mixture of I5-hydroxyacyl-ACPs of even numbered carbon chain 6'

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LAWRENCE I . GILBERT

length are formed. These range from C8 to C14.On further incubation with NADPH, enzyme and malonyl CoA, Wakil and his colleagues demonstrated the conversion of the &hydroxyacyI derivatives to their saturated homologues or palmitic acid. The final product was palmitic acid but in the presence of stoichiometric quantities of ACP, they were able to isolate palmityl-ACP. On the basis of these results they propose the following scheme for fatty acid synthesis:

+

(1) CH,COSCoA+ HSACP * CH3COSACP COASH (2) HOOCCHzCOSCoA HSACP HOOCCHZCOSACP COASH (3) CH3COSACP HOOCCHZCOSACP CH3CHOHCHZCOSACP COZ + HSACP (4) CH3COCHzCOSACP + NADPH + H + ---j CH3CHOHCHZCOSACP + NADP" (5) CH3CHOHCHzCOSACP * CH,CH=CHCOSACP Ha0 (6) CH3CH=CHCOSACP + NADPH + H * CHSCHZCHZCOSACP + NADP

+

+

+

+

+

+

+

Repetition of steps (3) to (5) six more times will yield palmityl-ACP which is then hydrolyzed to palmitic acid and ACP. There may be two systems for the biosynthesis of fatty acids. One is a mitochondrial system that may essentially be the reverse of &oxidation and is primarily concerned with the elongation of existing fatty acids by two carbon units (e.g. Cle to C18).The second pathway resides in the cytoplasm outside of the mitochondria and here one can demonstrate the conversion of acetyl CoA to palmitate (see Wakil, 1961, 1963 for an excellent review of this subject). This extra-mitochondria1system appears to function as in Fig. 14. Not all investigators agree that the main fatty acid biosynthetic system lies soleIy in the soluble fraction of the cell. Hulsmann (1962), for example, believes that mammalian heart sarcosomes incorporate acetate into long chain fatty acids by a mechanism similar to that of the soluble system. He goes so far as to suggest that the mitochondria are solely responsible for lipogenesis and that activity of the soluble system is due to mitochondrial enzymes that have escaped from their subcellular enclosures during fractionation. Lung tissue mitochondria also appear to be capable of synthesizing fatty acids (Tombropoulos, 1964). Thus, even in the more numerous experiments involving mammalian material, some doubt exists regarding the biosynthesis of fatty acids. As we shall see subsequently, information is sparse regarding the means by which insects synthesize unsaturated fatty acids and the situation is not greatly improved when we consider other animal forms

LIPID METABOLISM A N D F U N C T I O N I N INSECTS

129

(cf. Langdon and Philips, 1961). This problem is of significance to the insect since some insects require one or more unsaturated fatty acids in their diet (see Section D), and they constitute a large percentage of 0

II

CH3C-OH

\ / CoASHf ATP

CH~C-SC~A

CO,

+ ATP

3ADP +HP,O,+

O It C-OH

I

CHZ

I C-SCoA

0

ti

II

0

CH3C- SCoA

+ 2 NADPH +H+

CH,CH~CH,C--SCOA I

:a .G

C, Fatty acid

+CO,

FIG.14. Simplified scheme of fatty acid biosynthesis. a. Formation of acetyl CoA. b. Formation of malonyl CoA in the presence of COa through a biotin-COa complex. c. Condensation of several reactions whereby malonyl CoA acetyl CoA yield butyryl CoA COa. d. Butyryl CoA+malonyl CoA yields a Cg fatty acid through a series of reactions. Once rnalonyl CoA is present in sufficient quantities, acetyl CoA is needed only in the formation of the C4 fatty acid. After this, the process proceeds by the addition of rnalonyl CoA and the evolution of COa resulting in the lengthening of the molecule by one Ca unit at a time. For the formation of stearic acid the overall reaction would be: acetyl CoA 8 rnalonyl CoA 16 NADPH H stearyl CoA 16 NADP 8 COa 8 CoASH

+

+

+

+

+

+

-

+

+

+

the total fatty acids present in others (see Section B). Bloomfield and Bloch (1960) have demonstrated that anaerobic yeast possesses a mechanism for desaturation of palmitate to palmitoleic acid and stearate to oleate. In their view, the fatty acids are first converted to CoA thioesters and then desaturated by a mechanism involving formation

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LAWRENCE I . GILBERT

and dehydration of hydroxyl intermediates. If this mechanism applies to insects as well, then desaturation occurs subsequent to synthesis of the carbon chain. Perhaps, in those reports where insects incorporated labelled precursors into saturated fatty acids only, the experiments were not allowed to proceed for a long enough period of time. Mead (1963) suggests that the pathways of biosynthesis of monoenoic acids in animals may be quite different from those leading to saturated fatty acids. If this proves true, then we are at present completely ignorant of this phenomenon in insects. In a more recent investigation, Schroepfer and Bloch (1965) demonstrated the stereospecificity of the reaction responsible for the conversion of stearic to oleic acid in bacteria. The desaturating enzyme removes only one of each pair of hydrogens at Cg and Clo. The precise mechanism of desaturation is still unknown although it appears that hydrogen removal at C9 occurs prior to the corresponding desaturation at Clo. B . FATTY A C I D BIOSYNTHESIS I N INSECTS

That insects can synthesize long chain saturated fatty acids from simple precursors has been shown numerous times in various species (see for example, Bloch et al., 1956 and Rajalakshmi et al., 1963). In the course of these experiments that utilize labelled precursors, unsaturated fatty acids also exhibit the label. When the green peach aphid (Myzus persicae) is fed through an artificial membrane on a diet containing 18% sucrose solution with the addition of either [1-l4C]acetate or [U-14C]glucose,the insect was found to incorporate 75% of the label into stearic, palmitoleic and oleic acids (Strong, 1963b). Small quantities of myristic, linoleic and linolenic acids also contained label although there was no significant incorporation of these precursors into short chain fatty acids. It is of interest to note that both aseptically reared and non-aseptic adult boll weevils (Anthonornus grandis) synthesize long chain fatty acids from acetate at about the same rate (Lambremont, 1965). Palmitic, oleic and stearic were the major fatty acids synthesized but myristate, palmitoleate and oleate also were labelled when [14C]acetatewas injected. In addition to demonstrating the biosynthetic capacity of these insects, Lambremont’s results also suggest that the animal’s symbiotic flora make no significant contribution to fatty acid biosynthesis. The inability of Anthonornus to synthesize linoleate is in accordance with this fatty acid’s important role in the nutrition of several insects.

, ‘

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131

By extracting whole larvae and pupae of B. mori, Sridhara and Bhat (1964) studied the fate of [l-14C]acetate ingested by, or injected into fifth instar larvae. As one would expect, the more metabolically active larvae incorporated almost twice as much label into lipid as did the pupae, and the radioactivity in the saponiiiable fraction was double that in the unsaponifiable fraction. The major component of the latter mixture was hydrocarbon and it is possible that this was due to the synthesis of cuticular lipids since whole insects were extracted. The most heavily labelled fatty acids in this case were the saturated molecules, predominantly palmitate and stearate. The most heavily labelled unsaturated fatty acid was oleate and no label was detected in either linoleate or linolenate. It appears from this work and that on Prodenia by Zebe and McShan (1959) that saturated acids are synthesized at a higher rate in Lepidoptera than the unsaturated series. This contrasts with much of the work on the Orthoptera and Diptera, although Tietz’s (1961) in vitro studies suggest that palmitate is the major fatty acid synthesized by locust fat body. Recently, Bade (1964) demonstrated that after the injection of [l-14C]acetate,EurycotisJloridana showed a labelling pattern consistent with the classical view of the biosynthesis of fatty acids by the condensation of C2 units. Oleate composed about one half of the total newly synthesized fatty acids while palmitate was the most abundant saturated fatty acid. This is in agreement with the data of Sedee (1961) on Calliphora erythrocephala. Bade detected only traces of fatty acids having more than 18 carbons, and arachidonic acid was absent. From her data, Bade postulates that oleate is synthesized by desaturation of stearate. This is in accordance with the views of Bloomfield and Bloch (1958) but disagrees with Sedee’s (1961) supposition that insects use two different mechanisms for the synthesis of saturated and unsaturated fatty acids. Although this work of Bade’s was conducted in vivo rather than on individual tissues, it is significant because xenic and aseptic cultures were utilized. Louloudes et al. (1961) had demonstrated the synthesis of linolenate by another cockroach, Blatella, but this multiple desaturation may have been a consequence of gut symbionts. The main difference between the fatty acids of Eurycotis and those of higher animals is the virtual absence of fatty acids with a chain length greater than CI8 in the insect. Although the techniques of injection of labelled precursors, followed by the extraction of the whole insect (aseptic or non-aseptic) and radioassay of the resulting fatty acids, do tell us if the insect possesses the biosynthetic capability of producing fatty acids, they give us relatively

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little information regarding the pathway of synthesis, the enzymes involved or cofactors required. To elucidate these latter points, we must turn to in vitro studies. This is in no manner intended to deprecate in vivo experiments, for even while using homogenates or sub-cellular fractions, we must still strive to extrapolate the results back to the living insect. One of the pioneering in vitro studies on fatty acid biosynthesis in insects was that of Zebe and McShan (1959) on the fat body of the adult lepidopteran, Prodenia eridania. Acetate was readily incorporated into long chain fatty acids but required malonate, ATP, CoA and a reducing agent such as glutathione or cysteine (containing SH groups) for maximum incorporation. The major fatty acid produced was palmitic although small amounts of stearate, oleate, myristate and laurate were also identified. Homogenates had less than one-third the activity of intact fat body in synthesizing fatty acids. Zebe and McShan also demonstrated a slight conversion of [14C]glucoseinto fatty acid and showed that muscle was only one-fifth as effective as fat body in synthesizing fatty acids. In an elegant series of studies on the capacity of locust fat body homogenates and sub-cellular fractions to synthesize fatty acids, TietzDevir (1963) showed that homogenates were optimally effective when the incubation medium was supplemented with ATP, MgC12, glutathione, KHC03 and malonate. Addition of CoA and NADPH further stimulated the synthesis (Table XI). Although the use of citric acid cycle intermediatesdid not relieve the dependency of the system on malonate, they resulted in even greater stimulation when added in the presence of malonate. Of these intermediates, a-ketoglutarate was most effective. When the particulate matter was removed from the homogenate by centrifugation at 20,000 x g for 20 min, the supernatant proved to be as effective as the original homogenate. The newly formed fatty acids were esterified with glycerol almost immediately after synthesis and the FFA did not accumulate to any sigtllficant degree. Whereas in the homogenate 18% of the newly synthesized fatty acids were in the form of phospholipid and 8 1% as glycerides, in the supernatant 10% were in the phospholipid fraction and 90% were in the form of glycerides. Of the fatty acids synthesized, more than half was palmitate. The optimal conditions cited above were similar to those utilized in mammalian and avian systems (PopjAk and Tietz, 1954; Porter et al., 1957). The function of the malonate is not known but Tietz-Devir believes that the bicarbonate is utilized in the conversion of acetyl CoA to malonyl CoA. Kallen and Lowenstein (1962) had shown with non-

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133

insect material that stimulation of fatty acid synthesis by isocitrate and malonate is not linked to NADPH generation but is more likely connected with the synthesis of malonyl CoA. Using 14C02, Tietz-Devir also demonstrated CO, fixation by a combined particulate and supernatant system from the locust fat body. 91% of the label was recovered in malonate, 9% in acetate but none in the newly synthesized fatty acids. The particulate fraction was necessary for C02 fixation and for activation and decarboxylation of malonate. Thus in the supernatant fraction, malonate and bicarbonate TABLE XI Requirements for fatty acid synthesis by fat body homogenates of Locusta migratoria Cofactor omitted None1 ATP CoA Glutathione NAD NADP MgCL MnS04 Malonate a-Ketoglutarate KHC03

Acetate incorporated into fatty acid (pmoles) 0.33 0.05

0.14 0.17 0.33 0.19 0.01 0.32 0.01 0.20 0.08

Complete system contained (in pmoles); acetate (5); ATP (5); CoA (0.1 mg); Glutathione (5); NAD (0.5); NADP (0.5); MgCla (10); MnSOI (0.5); Malonate (20); a-Ketoglutarate (10); KHCOa (lo), and 0-6 ml (2 mg protein) homogenate in a total volume of 1 ml. pH 7-0. (From Tietz, 1961.)

stimulate fatty acid synthesis but yet the supernatant cannot metabolize these compounds. This contradiction is unexplained at present. The above, although by no means an inclusive survey, does support the proposition that many insects can synthesize -longchain fatty acids from acetate, presumably by the same means employed by both their inferiors (micro-organisms) and superiors (vertebrates) on the evolutionary scale. Once these FFA are synthesized, they are shunted into TGL and PL as can be seen by the relatively small quantities of FFA present. Since the biosynthetic pathway of the glycerides and phospho-

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L A W R E N C E I . GILBERT

lipids is in part a common one, we will discuss these two lipid types together. C . P H O S P H O L I P I D A N D TRIGLYCERIDE

1. General mechanism of synthesis

Again we must call on data derived from investigations of non-insect material to describe both the biosynthetic pathway of the glycerides and phospholipids as well as the function of PL. Figure 15, derived mainly Glycerol

ATp

a-Glycerophosphatef ATPI

Di-hydroxyacetone phosphate

NADH

1 2 Acyl-CoA

Phosphatidic acid /

Phosphatidyl inositol

Phosphatidylcholine

7

YCDP-choline

inositol

CDP-diglyceride

ie

a-Glycerophosphate

Phosphatidylglycerophosphate AcYI-COA

Phosphatidylglycerol CDP-diglyceride

riglyceride

Cardiolipin

FIG.15. The biosynthesis of phospholipids and triglycerides. (From Kennedy, 1961, 1963.)

from the classic experiments of Kennedy and his colleagues, reveals the close interrelationship between the synthesis of these two types of lipid. In addition to the pathway via a-glycerophosphate to TGL, another subordinate pathway via monoglyceride has been suggested (Hubscher, 1963). Monoglyceride

acetyl CoA

+ diglyceride

acetyl CoA

t

triglyceride

In this discussion we will lean heavily on the work of Kennedy’s laboratory and will allude to aspects of TGL synthesis as well (Kennedy,

LIPID METABOLISM A N D F U N C T I O N I N I N S E C T S

135

1961, 1963; Weiss and Kennedy, 1956; Weiss et al., 1960). For a complete review of the phospholipids, the reader should consult Ansell and Hawthorne (1964). It now appears that phosphatidic acid is the key compound in the synthesis of PL and TGL (Fig. 15). This compound is derived from a-glycerophosphatewhich itself results either from the phosphorylation of glycerol or reduction of dihydroxyacetone phosphate. It was at first believed that the glycerophosphate was mainly a consequence of the glycolysis of glucose. Recent data indicate the possibility of free glycerol acting as a precursor for the a-glycerophosphate through interaction with ATP (Isselbacher, 1965; Fig. 16). In at least some insects, there is a high glycerol titre in the hemolymph during the pupal stage and this becomes drastically reduced during adult development (Wyatt and Meyer, 1959; Wilhelm et al., 1961). It is possible that this glycerol may be used for the formation of a-glycerophosphate which then enters the pathway for the synthesis of PL or TGL, both of which are required for the development of the adult insect. Of importance is the requirement for the cytidine coenzymes. For the cell to obtain CDP-ethanolamine, ethanolamine must react with ATP to yield phosphorylethanolamine which in the presence of CTP gives rise to CDP-ethanolamine. Analogous reactions occur with choline finally to produce CDP-choline. Phosphatidic acid can be dephosphorylated in the presence of phosphatidic acid phosphatase to yield a D-a,&diglyceride which in turn can be converted to TGL, phosphatidylethanolamhe (PTE) or phosphatidylcholine (PTC). The DGL can also be rephosphorylated to phosphatidic acid by diglyceride kinase and in some instances this may be an important pathway (Hokin and Hokin, 1959, 1960; Strickland, 1962). Phosphatidic acid and the compounds derived from it contain fatty acids of varying degrees of unsaturation and of different chain lengths. As Kennedy (1963) suggests, these fatty acids bound to PL can be an important source of reserve energy for the cell. The scheme outlined above points out the close relationship between glycolysis, PL synthesis and TGL synthesis (see also Fig. 17). If for example, glycolysis was inhibited prior to the formation of pyruvate, there is a possibility that C3 units from glucose might be converted to lipid in the form of glycerides or PL. If there is an uninhibited flow from glucose to acetate, the acetate may give rise to fatty acids and join that moiety of the PL or glyceride molecule. It is now generally believed that the biosynthesis of choline and PL proceeds from the decarboxylation of serine and includes three methyla-

136

LAWRENCE I . G I L B E R T

CH,OH

I

CHOH

HOCH

I

0

CH,0P03H, 0 0

II

R-CH,C-OCH

II

II

2 R-CH,C-SCoA 2 CoASH

CH,OC-CH,-R

I I

CH,OPO,H,

0

II

CHZOC-CHZ-R

0

II

R-CH,C-0-CH

CH,&-CH,-R

I I

0

II

CH,OC-CH,-R FIG.16. Triglyceride biosynthesis from glycerol. a. Glycerol + ATP yields L-a-glycerophosphate in the presence of glycerokinase. b. a-Glycerophosphate+ 2 fatty acyl CoA derivatives yields phosphatidic acid. c. Phosphatidic acid is converted to a diglyceride in the presence of phosphatidic acid phosphatase. d. Diglyceride is converted to triglyceride.

137

LIPID METABOLISM A N D F U N C T I O N IN INSECTS

/Phospholipid

Glycerides

\

Carbohydrates

1 1

Phosphatidic acid COASH Sphingolipids *.-T Triosephosphate + a-Glycerophosphate+ ir Long-chain acyl-CoA2

t f

P

M

a

l

o

\Fatty acid

nyI-CoA

";" '.t\

Amino --f Acetyl-CoA A A c e t o a c e t y l - C o A *Methyl glutaryl-CoA acids J bcitrate Acetoacetate Terpenes Acetylations Oxalacetate 2 coz Sterols

3

J

\

+

FIO.17. Some interrelationships between lipid, carbohydrate and protein metabolism. (From Lynen et al., 1963.)

tion reactions (cf. Wagner and Folkers, 1964). Choline itself can be incorporated into PL in the mitochondrion while serine can be incorporated into phosphatidylserine prior to any decarboxylation. This latter reaction is thought to proceed in the microsomes. Thus, amino acids can become an integral part of PL synthesis.

2. General functions of phospholipids Phospholipids play a variety of roles in the life of the cell. Their presence in cell membranes has been unequivocally accepted. For example, the human erythrocyte membrane has about 59% PL of which about 35% is PTC and 25% PTE (Wolfe, 1964). The RNA-free microsomal fraction of neuronal membranes contains about 65% phosphatides. Petrushka et al. (1959) believe that PL is responsible for the spatial configuration of the mitochondrion, since these organelles rupture when PL is removed. Their low solubility in water makes the PL ideal substances for use in membranes that are vital for both the partitioning of separate cells and the microcompartmentalization of the cytoplasm. Their importance in oxidative phosphorylation is well known (Fleischer et al., 1961; Garbus et al., 1963; Green, 1962). Wojtczak et al. (1963) postulate that the phosphatidic acid formed during mitochondria1 contraction may be involved in tlfe mechanism of the active extrusion of water from the mitochondria. In fruits, it is

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LAWRENCE I . GILBERT

suggested that the initiating signal for senescence may be triggered by ethylene acting on the permeability of the mitochondria1 membrane (Biale, 1964) and may be affecting phospholipid in the membrane. A large amount of data has accumulated that links PL with transport of sodium ions (cf. Wolfe, 1964). A possible intermediate for this cationic transport is a labile phosphate attached to specific phosphatides or a lipo-phosphoprotein complex. This compound is believed to be rapidly phosphorylated by ATP through the activity of membrane ATP’ase. Subsequent dephosphorylation may lead to changes in the configuration of membrane molecules and this perhaps controls cation transport. The main proponents for the role of PL in active transport are L. and M. Hokin (Hokin and Hokin, 1955, 1959, 1960, 1961). Their original observations were on the effects of acetylcholine and various mammalian hormones on PL turnover in several tissues. A few of these substances increased the turnover rate of phosphatidic acid. In their view, ATP and DGL act at the interior of the cell membrane to form phosphatidic acid. The sodium salt of phosphatidic acid then diffuses through the membrane, is hydrolyzed on the exterior surface and results in the release of sodium and phosphate ions. In this way, sodium ions are actively transported. As Ansell and Hawthorne (1964) point out, much of the evidence for PL involvement in the sodium pump is circumstantial and there are many valid objections to the theory. However, it is still an open question. The adult mammalian brain contains between 20-25% of its dry weight as PL. With all the numerous papers published on the analysis and metabolism of PL in the brains of higher animals “no precise function has been attributed to any single component” (Ansell and Hawthorne, 1964; p. 278). Notwithstanding the absence of precise information, PL have been implicated in ageing of the nervous system, regeneration and in various diseases affecting the nervous system. The insect nervous system contains quantities of PL of the same order of magnitude as those reported for other animals (Heslop and Ray, 1961). The lipids of the bee brain for example, contain PTE (12% of total lipid), PTC (2%) and sphingomyelin (1.5%) (Patterson et al., 1945). It is thus probable that PL may play a comparable role in the nervous integration of insects as they do in higher animals. Beef heart mitochondria contain about one fourth of their weight as lipid, of which 90% is PL (the major constituents being PTC and PTE). Biochemical and electron microscopic studies leave no doubt of the central role played by lipids in electron transfer and oxidative phosphorylation within the mitochondrion (Green and Fleischer, 1963). In

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insects for example, NADH-cytochrome c reductase, cytochrome b, and cytochrome oxidase are for the most part located in the “lipid rich particle fraction” of centrifuged homogenates of Bombyx embryos (Chino, 1963). The mitochondria lose much of their biochemical activity after acetone extraction but this can be restored by the addition of coenzyme Q (CoQ) (see Section D) and PL. Green and Fleischer now visualize the mitochondrion as having a PL requirement at three points along the electron transfer chain : succinate

Q:QHa

-

cytochrome c:cytochrome c reduced

-

O2

They suggest that PL interacts with the mitochondria1 proteins in two types of reactions. One is ionic between the acidic PL and basic proteins like cytochrome c while the other is hydrophobic between the PL and structural protein. One of the properties of the PL which make them indispensable to the mitochondrion is the fact that they orient in water to form micelles. The amphipathic character of the PL molecule allows it to act as a bridge between water and protein. Green and Fleischer propose that the PL provides a non-aqueous medium (giving it a low dielectric constant) in which specific reactions can take place. The oxidationreduction reactions along the electron transfer chain appear to be oriented in such a way so as to be in contact with or actually lay within the PL layer that envelops the complex. Without PL, the mitochondrion ceases to function. In addition to the PL and ubiqcinone content of mitochondria, lipids may be important in another aspect of mitochondria1 function. Lewis and Fowler (1960) showed that in blow-fly flight muscle mitochondria, an endogenous lipid uncoupling agent was generated enzymatically. This had previously been suggested from the work of Lehninger and Remmert (1959) in rat liver mitochondria. It is proposed that this lipid may be a crucial factor in the reversible swelling and contracting of mitochondria; a phenomenon that may be vital to mitochondria1 regulation. 3. Metabolism and function in insects Having denoted some of the functions of PL we can now turn to PL and TGL in insects. Little can be tabulated regarding the synthesis of TGL in insects other than what we have discussed previously. No work has been reported on this biosynthetic pathway in insects although we know the fat body readily esterifies FFA intu TGL (Section 11). The elucidation of the role of the cytidine cofactors in the insect would

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be of real interest but little has appeared on this subject as of this time. In a histochemical study of the larval fat body of the dermestid beetle, Anthrenus uorax, Nair and George (1964) showed that the peripheral globules of the fat body cells are composed of a PL-protein complex, whereas the centrally located globules are composed of glycerides. On the basis of this microscopical study, they propose that the peripheral globules are lipopoietic centres where dietary amino acids are deaminated and converted into specific lipids. The lipids are then supposed to be transported to the centre storage globules. Although an interesting hypothesis, there is little or no biochemical evidence for or against it. It is at best, difficult to draw conclusions regarding the temporal sequence of a biochemical process on the basis of microscopical observations. As mentioned previously, several insects have large concentrations of glycerol in their hemolymph (Wyatt and Kalf, 1957; Wyatt and Meyer, 1959; Salt, 1957; Wilhelm et al., 1961). Salt (1959) has shown that larvae of the sawfly Bracon sephis contain concentrations of glycerol as high as 25% (5 molal) in the hemolymph. In H . cecropia, glycerol may constitute up to 3.5% of the hemolymph and tissues of diapausing pupae. This glycerol appears to be derived from glycogen. In diapausing embryos of B. mori, both glycerol and sorbitol are present (Chino, 1958). It is possible that this glycerol may be a precursor of PL and TGL in these insects in addition to acting as an “anti-freeze” (see Touster and Shaw (1962) for a review of the biochemistry of polyhydric alcohols). Our results (Habibulla and Gilbert, 1965) indicate a high rate of incorporation of labelled glycerol into the phospholipids of various tissues of developing adults of H. cecropia. As part of a survey of invertebrate phospholipids, Hack et al. (1962) studied insects representative of five orders. All contained PTE, PTC, monophosphoinositide, polyglycerophosphatide, ethanolamine plasmalogen and traces of choline plasmalogen. Thus, insects appear to contain the same general types of PL as the vertebrates. A great deal of the work on PL in Lepidoptera has been conducted on Galleria mellonella with a view towards explaining the mechanism of wax digestion. Galleria larvae possess a remarkably high concentration of PL in the hemolymph (about 22% of the hemolymph lipid) and on a percentage basis contain twice the amount present in mammalian plasma (Wlodawer and Wisniewska, 1965). There appears to be a discrepancy regarding the PL constitution of waxmoth larval hemolymph with Lenartowicz (1961 ; Lenartowicz and Niemierko, 1964;

141 Lenartowicz et al., 1964) demonstrating large quantities of phosphorylethanolamine and phosphorylglycerol in the larval lipids, and more phosphorylethanolamine than phosphorylcholine in the hemolymph. These investigators assert that the concentration of phosphorylcholine in the hemolymph is the highest reported for any animals. Both the phosphorylcholine and phosphorylethanolamine concentrations exceed those reported for B. mori, H. cecropia, A . polyphemus and P. sexta (Wyatt et al., 1963). On the other hand, Wlodawer and Wisniewska (1965) maintain that PTC makes up 60% of the hemolymph PL while PTE constitutes only 20%. Sphingomyelin was not detected. The possibility exists however that phosphorylcholine which is a precursor of CDP-choline and PTC, is present in higher quantities due to a block leading to PTC, while the titre of phosphorylethanolamine remains low because it is rapidly converted to PTE. This is speculation however, and the discrepancy has not yet been resolved. The means by which these compounds come to be in the hemolymph in such great quantities has not been adequately explained. In their studies of H. cecropia, Wyatt’s laboratory (Carey and Wyatt, 1963; Wyatt et al., 1963) showed that the concentrations of phosphorylcholine and phosphorylethanolamineare higher in the hemolymph than in the fat body and the enzymes necessary for the synthesis of these PL appear to be absent from the hemolymph. With this circumstantial evidence, they suggest a selective secretion of these compounds from the fat body into the hemolymph. The role of PL in the hemolymph is obscure but they may be involved in lipid transport as they are in mammals (see Section 11). Our data on H. cecropia (Habibulla and Gilbert, 1965) reveal that although PTC and PTE are the predominant PL of the fat body and other tissues, phosphatidic acid is the major PL component in the pupal hemolymph. In fact, phosphatidic acid and cardiolipin constitute 80% of the pupal hemolymph PL but this decreases during adult development to reach a low of 3%. The hemolymph phosphatidic acid is utilized during development as a precursor for TGL and PL synthesis in the fat body. It is of interest in this regard that histochemical data have implicated the oenocytes as the major depot for PL in the locust rather than the fat body cells (Coupland, 1957). In waxmoth larvae, the hemolymph is suggested to be a phospholipid “depot” (Lenartowicz and Niemierko, 1964). This would have no counterpart in higher animals but Galleria appears to be an atypical animal on the basis of its diet. These same authors suggest that phosphorylethanolamine is utilized to a higher degree than”its choline counterpart during starvation and that its metabolism is independent of L I P I D METABOLISM A N D FUNCTION I N INSECTS

142

LAWRENCE I . G I L B E R T

phosphorylcholine. On the other hand, Wlodawer and Wisniewska (1965) found little qualitative change during the period of food deprivation. The question remains open. In addition to the PL discussed above, some Lepidoptera contain a minor component containing alanine but its nature has not yet been resolved (Hodgson, 1965). In a developmental study of the PL content of B. mori, Niemierko et al. (1956) showed that the quantity of PL as percentage of total lipid reaches a maximum at the end of metamorphosis. They postulate synthesis of PL toward the latter part of adult development. Our work on Cecropia supports this contention and suggests that it is due mainly to the morphogenesis of the giant mitochondria in the developing flight muscle (Habibulla and Gilbert, 1965). We must take care in evaluating differences between synthesis and turnover. For example, the fatty acid portion of PTC is known to turn over without either synthesis or breakdown of the remainder of the molecule (Lands, 1958). Thus incorporation of label into PL does not necessarily imply synthesis. Only quantitative evaluation by gravimetric procedures will allow a true differentiation between these two alternatives. The study of Carey and Wyatt (1963) suggests a steady turnover of PL in the Cecropia fat body. Radiotracer studies also indicate a rapid turnover of hemolymph PL but this in turn is most likely due to turnover within other tissues and subsequent release into the hemolymph. In contrast to the results cited above on Bombyx, Sridhara and Bhat (1965a) reported that 23% of the larval lipid of this insect is PL while the lipid of pupae and moths is only 18 to 19% PL. Of the various larval body parts analysed, silk gland had the highest concentration of PL (68% of total lipid) whereas about half the total lipid extracted from the hemolymph and intestine was PL and only 3% of the fat body lipid was PL. Of interest is the finding by these investigators (1962) that the PL of Bornbyx contains about 20 times more choline than ethanolamine. This is in contrast to the PL constitution reported for Diptera and Orthoptera which will be discussed subsequently. In addition, a considerable amount of sphingosine was found whereas it is absent from the Diptera. These results are similar to those of Chojnacki (1961; Chojnacki and Korzybski, 1962, 1963) working on another moth Acta caia. In the adult there is about twice as much PTC as PTE with a relatively high concentration of sphingomyelin. It appears that these PL in Acta are utilized during adult life as the quantity of lipid phosphorus diminishes with the age of the moth. Using 32Porthophosphate, these workers demonstrated a higher rate of incorporation into PTE of Celerio eurphorbiae adults than into the PTC. On the basis

LIPID METABOLISM A N D FUNCTION I N INSECTS

143

of specific activities, they suggest that PTC is incorporated in toto into PTE. According to present information however (Fig. 15) it is more likely that phosphorylcholine is first converted to CDP-choline which combines with diglyceride to yield PTC. Investigations of the PL of Diptera have been primarily concerned with the analysis and incorporation of nutrient precursors into these compounds. In contrast to higher animals, the major PL fraction has been identified as PTE (Bieber et al., 1961) and in A . aegypti this PL comprises 86% of the total extractable PL (Fast and Brown, 1962). Bieber et al. (1961) demonstrated that when larvae of Phormia regina were reared on a carnitine containing medium, a new “lecithin” was recovered. This compound constituted 20% of the total PL and was eluted prior to PTC on silicic acid columns. It contained 8-methylcholine rather than choline and was absent from larvae raised on a choline containing medium rather than on one having carnitine. It is thought that the larvae have the ability to decarboxylate carnitine, and under conditions where choline is limited, other quaternary ammonia compounds can be utilized for structural purposes, such as the manufacture of membranes (Bieber ef al., 1963; see however, Agarwal and HOUX,1964). In 1963, Bieber and Newburgh demonstrated that dimethylaminoethanol and dimethylaminoisopropanol are incorporated into the PL of Phormia when they replace choline in the diet. Under these conditions the quantity of choline detected in the PL fraction was drastically reduced. It appears then, that the quality of the PL is not of great importance to the functioning of the organism and agrees with the contention of Fleischer et al. (1962) that although the respiratory activity of mammalian mitochondria depends upon the presence of lipids, it is not dependent on one specific lipid organization. Following the work on Phormia, it was found that Musca domestica larvae bred on a chemically defined diet under aseptic conditions could also incorporate 8-methylcholine into PL when the dietary source of choline is replaced by either carnitine or 8-methylcholine (Bridges et al., 1965). Although a low level of PTC was present in larvae grown on this choline-deficient diet, it was absent from the adult flies. It may be that the ability to decarboxylate carnitine is lost during metamorphosis. If this is truly a juvenile characteristic, it may be a proper mechanism for the study of the effect of insect hormones on the synthesis or activation of specfic enzymes or enzyme systems. Though grown on a diet containing carnitine, these flies were unable to synthesize phosphatidyl carnitine (see also Agarwal and Houx, 1964). In none of the studies discussed above was the function of PL in

144 LAWRENCE I . GILBERT insects dealt with. They were nutritional studies which showed that the insect was capable of certain biochemical feats. In general, the extraction of whole insects is not the most efficient method of studying physiological mechanisms. In an attempt to use specific sub-cellular organelles, Crone (1964; see also Price and Lewis, 1959) analysed the PL content of the flight muscle sarcosomes of M. domestica. The sarcosomal lipid phosphorus comprises about 7% of the total lipid phosphorus in the adult female or about 210 mg of PL/g of protein. This figure is close to that reported for beef heart mitochondria (Fleischer et al., 1962). Relatively, the proportion of total sarcosomal PL existing as PTC is much lower than that of the whole fly (Crone and Bridges, 1963). After injection of 32P,the PTC was shown to have a greater specific activity than the PTE. Of the total lipid phosphorus in the sarcosomes, 69% was due to PTE; 9% to PTC and 6% to polyglycerolphosphatide. Most of the polyglycerolphosphatide of the fly was located in the thorax and as in mammalian tissues, was associated with the mitochondria. It is most likely that PL in insect mitochondria play a role similar to that played in mammalian mitochondria. The function of PL in the insect sarcosome is certainly an open question and one well worth answering. In addition to their most certain role in the post-embryonic development of insects, PL play an important role in insect embryogenesis. The egg lipid of L. migratoriu, for example, is composed of about 20% PL, which actually constitutes about 5% of the total dry weight of the egg (Allais et al., 1964). PTE predominates but phosphatidylserine, PTC and a sphingomyelin-like lipid were also identified. There was little variation in PL during embryogenesis although blastokinesis occurred simultaneously with an 11% decrease in PTC. Perhaps the energy for this vital embryonic process was supplied by PTC. Allais and his colleagues also demonstrated synthesis of PL at the termination of blastokinesis. This phenomenon of PL synthesis during embryonic development has also been reported for Bombyx (Niemierko et al., 1956). It is likely that as cell differentiation progresses more PL is needed for the construction of cellular and sub-cellular membranes. Insects then, can synthesize many of the long-chain fatty acids and esterify these into PL and TGL. Fatty acid biosynthesis is most likely a process very similar to that found in other organisms. The data regarding the synthesis of PL and TGL are sparse but until contrary evidence is forthcoming, we must postulate that here again, the insect does not break biochemical regulations. Can insects, however, synthesize all the fatty acids necessary for life?

L I P I D METABOLISM A N D F U N C T I O N I N INSECTS

145

D. FATTY ACIDS I N NUTRITION

Although oleic acid can be synthesized by vertebrates, linoleate and linolenate are thought to be dietary requirements. Linoleate in fact is believed to be a primary precursor for essential fatty acid biosynthesis in mammals (cf. Wagner and Folkers, 1964). A fatty acid deficiency in mammals is characterized by low survival rate, subnormal growth, scaly skin, skin lesions and impaired ability to reproduce. One can draw a close analogy between these nutritional defect symptoms and those reported in insects. For reviews of insect nutrition see Friend (1958) and House (1962, 1965). More than twenty years have passed since Fraenkel and Blewett (1945, 1946) demonstrated that wheat germ oil was needed for the normal development of Ephestia kuhniella. The critical substance in the wheat germ oil was identified as linoleic acid. This unsaturated fatty acid is needed for the normal formation of adult scales and is perhaps necessary for either the production of moulting fluid or the proper functioning of the moulting fluid. Linolenate was equally effective but oleate was an inadequate substitute, indicating that this insect cannot convert oleate to linoleate. In 1947, these same investigators showed that Tenebrio larvae do not require linoleate, indicating that linoleate or linolenate can be synthesized by Tenebrio (or its symbionts). Since this early work there have been numerous reports on the necessity in the diet of various fatty acids if the normal physiological processes of the insect are to continue. For example, Gordon (1959) demonstrated that the progeny of Blattella germanica that were raised on a linoleate deficient diet showed deficiency symptoms (see also Pant and Pant, 1961). Dadd (1960a, b) corroborated the need for linoleate in L. rnigratoria and S. gregaria. Although larval growth of GalZeria is not affected by a diet devoid of linoleate, successful emergence of the adults from the pupal cuticle depends on the presence of linoleate or linolenate in the diet (Dadd, 1964; see also Vanderzant et al., 1957). In Argyrotaenia velutinana, linoleic or linolenic acids are needed for normal growth. Gas chromatographic analysis revealed that pupae of this species grown on deficient diets did not contain either linoleate or linolenate (Rock et al., 1965) indicating a biosynthetic block. From the vast nutritional literature one can conclude that either several insects cannot synthesize linoleate or linolenate from less complex precursors and so are similar to mammals, or the proper precursors are not yet known and are thus omitted from the defined diets. However, the questions as to how these fatty acids promote normal growth and where the metabolic blocks exist, are still

146

LAWRENCE I. GILBERT

unanswered. While on the question of fatty acids and nutrition, it is of interest to note that in the locust, the maternal diet influences the lipid content of young locusts (Blackith and Howden, 1961); the reasons for this are unknown. One problem regarding this nutritional work concerns the purity of the fatty acids. Until fatty acid analyses could easily be conducted with gas chromatography, the investigator had to trust his commercial source. In our experience, many of the so-called pure lipids produced commercially are grossly contaminated and perhaps these contaminants have a nutritional value. The data of Chippendale et al. (1964) support this contention. Their work on the dietary requirements of the cabbage looper, Trichoplusia ni, suggest that linolenate is an essential nutrient and that linoleate cannot replace it. This is the reverse of the situation in the rat and many of the insect studies. These investigators used fatty acids of extremely high purity and point out that many of the past effects attributed to linoleate may have been the responsibility of contaminating linolenate. This infers a critical re-evaluation of all the past work where commercial fatty acids not analysed by the investigator were used. As Dadd (1963) points out, the dietary requirements for unsaturated fatty acids that have been demonstrated, were demonstrated on only a few species of Lepidoptera and Orthoptera. Thus we are led to the possibility that some insects may indeed be able to synthesize all required fatty acids. The lipid content of the insect’s diet may have physiological consequences far removed from those cited above. By increasing the lipid content of the larval diet of the pink bollworm, the insect can be induced to enter diapause at a greater rate (Bull and Adkisson, 1962). This finding is in agreement with those of Squire (1940) and Vanderzant and Reiser (1956). Since diapause is usually thought of as a hormonal deficiency (cf. Gilbert, 1964), the presence of a critical amount of lipid in the digestive tract, hemolymph or fat body may influence the synthesis and/or release of a hormone (perhaps through nervous stimuli acting on the neurosecretory cells of the brain). The large lipid store is of course important to diapausing insects as a ready energy source for this non-feeding stage. As we shall see subsequently, several of the insect hormones are lipoidal and lipid appears to be the precursor for their synthesis. In addition to possessing a growth promoting effect, fatty acids may also have a growth inhibiting effect in insects. A recent report claims that particular fatty acids can prevent the normal growth and development of young larvae of the orthopteran, Gryllodessigillatus (McFarlane

147 and Henneberry, 1965). In these experiments, methyl oleate and methyl palmitate were most effective. Curiously, they were only effective when applied to the exterior of the insect, presumably since this method of application bypassed any detoxication mechanism that may be present in the gut. Although the effects may be real, their significance remains obscure since one does not usually find insects wading through pools of fatty acid esters. As far as this author is aware, no data exist that conclusively demonstrates any nutritional requirements for the fat soluble vitamins. Perhaps any such requirements are met by the synthetic capabilities of symbiotic micro-organisms (see Henry, 1962 for a discussion of micro-organisms and insect nutrition). Notwithstanding the criticisms cited above, it may be that dietary fatty acids will be ultimately shown to have profound effects on insect growth. This is certainly the case for sterols (see Section VI), but insects cannot synthesize any sterol. LIPID METABOLISM A N D FUNCTION I N INSECTS

E. SUBSTRATE INTERCONVERSION

A final topic in connection with the build-up of lipid stores in insects is that of substrate interconversion. Not only are insects capable of utilizing ingested lipid and synthesizing fatty acids from acetate, but like most animals they are capable of converting nonlipoidal material into lipid. Indeed, in some insects this may in fact be the main source of body lipid. One of the most striking examples of the interconversion of carbohydrate to lipid in the animal kingdom, is the involuting mammary gland of the rat in which almost 100% of [14C]glucoseadministered, is recovered in the lipid (McLean, 1964). In this case the C2 units for lipid synthesis arise exclusively from glycogen. In lung tissue, glucose is converted to fatty acids and PL (Tombropoulos, 1964) while glucose is the precursor for glycerol in the mammary gland, and the glycerol is then used for the synthesis of milk lipids (Popjhk et al., 1952; Luick and Kleiber, 1961). In fact, the hydrogen atoms of glucose are utilized in the synthesis of mammary gland fatty acids by a pathway distinct from that which leads to the incorporation of the carbon atoms (Levy, 1963). The conversion of amino acids to lipid in mammalian systems is well known (see for example Feller and Feist, 1959). Figure 16 summarizes some of these general relationships. Although insects are devoid of mammary glands, it has been known for many years that they can convert non-lipid substances to lipid. Many of the earlier experiments consisted of feeding an insect a nonlipid diet and noting the increase in lipid with time by either extraction

148

LAWRENCE I . GILBERT

procedures or histochemical techniques (Abderhalden, 1925;Hofmann, 1872, MacCay, 1938; Melampy and Maynard, 1937; Mishikata, 1922; Weinland, 1908). For example, when mosquitoes were depleted of their lipid reserve by food deprivation and subsequently fed on a diet containing protein, amino acids and glucose, lipid reappeared in the fat body suggesting conversion of one or more of the above mentioned substrates into lipid (Wigglesworth, 1942). At pupation, B. mori contained twice as much lipid as could be accounted for by the leaves it had eaten, indicating conversion of other substances to lipid (Manunta, 1935; Uvarov, 1928). Another technique utilized was manometry where the RQ of the insect indirectly suggested the conversion of carbohydrates to lipid (Fulton and Chamberlain, 1934; Johannson, 1920; Timon-David, 1927). More recent investigations utilizing modern biochemical methodology support the proposition presented above. In an excellent in iiitro study of the locust fat body, Clements (1959) found that acetate, glycine, leucine and glucose are in part converted to lipid. Using labelled precursors, he showed that acetate was the most efficient lipid precursor and that less 14C02was recovered with acetate. The carbon dioxide was the result of the acetate passing through the citric acid cycle (Hines and Smith, 1963). A most interesting series of experiments on the mosquito Aedes was reported by Van Handel and Lum (1961). The female mosquito can be starved until virtually no TGL remains in the body but resumes neutral lipid synthesis when fed glucose. In contrast to the male mosquito or both sexes of housefly, the female mosquito could readily synthesize TGL when fed only glucose. The lipid synthesizing system of Aedes females is so efficient that the quantity of TGL may in some cases exceed the lean dry weight. Of the fatty acids identified in these TGL synthesized from glucose, palmitate, palmitoleate and oleate constitute 92% of the total. Polyunsaturated fatty acids were not synthesized under the experimental conditions utilized. These experiments dramatize the fact that at least some insects have a profound capacity to convert carbohydrate to lipid. We (Chino and Gilbert, 196%) have studied the interconversion of carbohydrate and fatty acid in Cecropia pupae and developing adults using [14C]labelled glucose, acetate and protein-bound palmitate. As seen in Table XI, when [14C]glucosewas injected, considerable radioactivity was recovered in the glycogen fraction in both chilled pupae and developing adults. A significant amount of radioactivity was also demonstrable in the glyceride fraction and as much as 2-2-5% of the label was recovered in glycerides extracted from developing pupae.

TABLE XI1

z z U

The fate of (U-14C)glucose injected into pupae a n d developing adults of H. cecropia Glycerides Stage

Control (chilled Pupa)

Free fatty acid

A

I

Sex

Male Female

Chilled Female Female (3-5

Female

co,

Glycogen A

>&r

Amount % ctslmin % ctslmin % Amount % recovery recovery (mg) fresh (mg) fresh wt. Wt.

s

I -

ctslmin

%

ctslrnin

recovery

%

m

recovery

0

479 471

-

113 115

0.01 0.01

8 7

0.00 0.00

78 65

-

2,550 2,700

0.25 0.27

-

-

364 366 291 342 332 349 341 249 334 349 316 273

7.8 7.7 6.5 5.7

2,410 2,115 4,250 3,006 23,760 21,055 17,830 17,840 22,9W 19,840 25,500 23,900

0.24 0.21 0.43 0.30 2.4 2.1 1.8 1.8 2.3 2.0 2.6 2.4

162 150 200 242 303 382 328 249 194 270 348 273

0.02 0.02 0.02 0.02 0.03

101 49 31 57 48 40 87

2.2 1.0 0.5 0.9 1-2

120,800 75,400 118,040 107,400 52,550 33,000 64,300 34,700 4,730 13,730 78,780 58,900

12.1 7.5 11.8 10.7 5.2 3.3 6.4 3.4 0.47 1.4 7.9 5.9

47,470 51,620 37,460 36,550 122,530 142,750 108,490 109,900 335,180 239,980 300,330 230,950

4.7 5.2 3-7 3.7 12.3 14.3 10.8 11.0 33.5 24.0 30-0 23.1

8.0

8.1 5.4 5.9 8.6 8.4 4.7 4.2

0.04

0.03 0.02 0.02 0.03 0.03 0-03

44 2 4 30 45

0.9 1.6 1.0 0.04 0.08 0.05 0.8

2 W

t:

-

~

E

~-

From Chino and Gilbert (1965~). Animals were sacrificed one day after injection of loEctslmin. The control consisted of injecting the ['*C]glucose into a pupa and sacrificing the animal immediately by freezing in a bath of dry ice and acetone. These animals were then analysed in an identical manner to the experimentals.

2 2 cl

=] 2

5

.-cl 2 CI

P

W

150

LAWRENCE I . GILBERT

The radioactivity found in the long chain FFA fraction was low but significant. Although a considerable amount of the label from [“CIpalmitate was incorporated into glycerides, we could detect no sign%cant incorporation into glycogen (Table XIII). Similar results were obtained with labelled acetate; that is, high incorporation into lipid, a large quantity combusted to C02, but no appreciable amount found in carbohydrate. These results support our previous contention that when FFA are synthesized they are readily incorporated into glycerides ; but more relevant to the present section is the demonstration that Cecropia can convert carbohydrate to lipid, but not the reverse. There have been several suggestions that insects possess the mechanism for converting lipid to carbohydrate, but for the most part this evidence has been circumstantial or only suggestive (Nair and George, 1964; Hitchcock and Haub, 1941 ; Levinson and Silverman, 1954). As far as this author is aware, the conversion of lipid to carbohydrate has not been unequivocally demonstrated in any metazoan although it is a reality in plants and certain microorganisms. In a careful study of the larval-pupal moult in Cecropia, Bade and Wyatt (1962) found that it was unnecessary to invoke a lipid to carbohydrate conversion to explain the changes in nutrient reserves during pupation, although by examining the changes in substrate one may have assumed such a conversion. It appears that the seeming increase in carbohydrate can be explained by a transfer of material from the cuticle to the interior. In fact, the glyoxylate cycle could not be demonstrated in Cecropia and this is prerequisite for such a substrate interconversion (Bade, 1962). It should be noted however, that isocitric lyase activity has been detected in pre-pupae and young pupae of Prodenia (Carpenter and Jaworski, 1962). According to these authors, the enzyme which converts isocitrate to succinate and glyoxylate is most likely maximally active at those developmental stages when the insect is consuming its lipid store. It may be that the past failure to demonstrate this enzyme in insect material is due to utilization of the wrong stage in the life history of the insect. The complete glyoxylate cycle still remains to be demonstrated in insects. Taking a teleological view, one can find a number of reasons why an insect, or any animal for that matter, would find it more beneficial to convert carbohydrate to lipid rather than the converse. The reasons have been stated by many and include the fact that lipid contains more energy per unit weight and yields more metabolic water than carbohydrate. Only if the insect lives in an environment of low oxygen tension or is dependent on anaerobic metabolism, would carbohydrate be

+?

21

7

Irr

r

P

U

TABLEXI11

z1

The fate of (l-14C)palmitate injected into pupae a n d developing adults of H. cecropia Glycerides Stage

Sex

Control (chilled pupa) Chilled pupa

Developing adult (3-5 days) Developing adult (1517 days)

Male Female Male Male Female Female Male Male Female Female Male Male Female Female

i

I {

From Chino and Gilbert (196%).

Amount % (mg) fresh wt. 403 336 386 371 298 333 349 439 274 247 345 356 201 232

8.7 8.0 5.5

5.8 7.1 8.6 4.8 4.7 8.8 9.2 4.3 4.2

ctslmin

)

% recovery

2,635 1,805 346,400 345,000 277,500 323,000 352,700 377,500 379,000 379,800 278,100 382,300 467,000 322,300

coz

Glycogen

A

i

U

0.03 0.02 34.6 34.5 27.8 32.3 35.3 37.8 39.9 38.0 27.8 38.2 46-7 32.2

zm

A

I

Amount % (mg) fresh wt. 85 98 76 84 85 120 55

107 60 59 0 0 21 53

-

1.7 1.8 1.5

2.0 1.4 2.0 1.0 1.1 0.0 0.0 0.45 1.0

cts/min

%

j-2

recovery 1,824 1,955 1,210 1,025 850 1,170 525 1,320 845 810 350 645

0.18 0.19 0.12 0.10 0.09 0.12 0.05 0.13 0.08 0.08 0.03 0.06

recovery 0

!2

89,930 76,620 . 55,230 76,090 56,790 63,170 61,790 80,720 194,000 148,900 226,400 138,900

-

-

8.9 7.7 5.5 7.6 5.7 6.3 6.2 8.1 19.4 14.9 22.6 13.9

E 9

1:

z

2 cI

2: c1

1: 0 +I vl

152

LAWRENCE I . GILBERT

preferred. Glycolysis, although inefficient, will yield enough energy for life processes to continue up to a point, but the oxidation of fatty acids will not occur in the absence of oxygen. Perhaps an intensive investigation of insects inhabiting an ecological niche where the pOz is low would uncover instances of a complete glyoxylate cycle and demonstrate lipid to carbohydrate conversion. Until such data are forthcoming, we must assume that this conversion does not take place in the Insecta. Thus far we have not discussed waxes, hydrocarbons, or the so-called cuticular lipids. These compounds do not really fit into the topic under consideration but are still lipids and are vital to the existence of the insect.

V. HYDROCARBONS A N D WAXES A . CUTICLE

It is now taken for granted that one or more layer of lipid on the exterior surface of the cuticle is in part responsible for the resistance of the insect to desiccation. This relationship between the cuticular lipid and its role in minimizing transpiration was demonstrated more than thirty years ago (Ramsay, 1935). In his review of the insect cuticle, Beament (1964) notes that “organized lipid plays the most important role in the passive regulation of water movement and in modifying the adhesion of water to the surface of cuticular membranes” (p. 68). According to Beament, the physico-chemical properties of the cuticular lipid layer suggest that the lipid acts as a “valve” or “rectifier”. He visualizes the organization of the lipid of the cockroach (P. americana) cuticle as consisting of a tightly packed single monolayer of lipid molecules up against the cuticular surface. Overlaying this monolayer is a “grease” twenty to thirty times as thick as the monolayer but arranged in a random manner. It is the monolayer, however, that is more impermeable to water. This arrangement probably varies in different insects but little research effort has been dedicated to the problem in insects other than cockroaches. In 1955, Beament suggested that shorter chain molecules may act as solvents and promote the tightly packed arrangement of the waterproofing layer. Unfortunately, very little is known of the biochemistry of this lipid, although it is a most promising area of investigation. There have however, been several chemical analyses of insect cuticular lipids and some may shed light on the mechanism by which this material is synthesized. The total hydrocarbon content of M . domestica is composed of unsaturated, cyclic and straight chain compounds (Louloudes

L I P I D METABOLISM A N D FUNCTION I N INSECTS

153

et al., 1962). These range in chain length from cl6 to CS5and are composed of both odd and even numbers of carbons although the odd numbered compounds predominate. This appears to be true for the Mormon cricket (Anabrus simplex) as well (Baker et al., 1960). The cuticular hydrocarbons of Musca differed from the total body hydrocarbons by having a higher concentration of lower molecular weight compounds. This finding appeared to corroborate Beament’s (1955) suggestion regarding the presence of “solvents ” in cuticular lipid. In 1963, Gilby and Cox reported on their analysis of the wax of the cast cuticle of P . americana (the same insect investigated by Beament). By utilizing column and gas chromatography as well as infra-red spectrometry, they demonstrated that hydrocarbons compose 75% of the cuticular wax. Fatty acids, esters and aliphatic aldehydes are present in roughly equal amounts and together with a trace of sterol account for the remaining 25%. The major component was identified as a C2, unconjugated diene, probably heptacosa-9,18-diene (see also Baker et al., 1963). The largest percentages of saturated hydrocarbons were C25and c 2 6 although most of the hydrocarbons were unsaturated. Table XIV reveals that the main fatty acids present are c16 and C18 while the predominant aldehydes are C14 and CI6. Of great interest is the absence of alcohols, except for sterols that comprise less than 1% of the ‘total wax. This finding negates the argument that the cuticular lipids are similar to other waxes. The results are remarkably similar to those reported for the Mormon cricket (Baker et al., 1960). The main points of difference are that the cricket wax contained more varieties of hydrocarbons and these were saturated; the cricket wax contained polyunsaturated C18 fatty acids whereas they are absent from the cockroach wax; the cricket wax was devoid of aldehydes but contained polymeric resins. As Gilby and Cox suggest, it is possible that these cricket resins are derived from aldehydic precursors. In 1962, Gilby reported the absence of natural volatile substances in the cockroach wax and suggests (Gilby and Cox, 1963) that the fluidity of the “grease” can be accounted for by involatile liquid constituents of the hydrocarbons and free acid fractions. Although the cuticular wax hardens when removed from the cuticle, it remains fluid in vivo. The Australian investigators propose that in vivo, an inhibitor prevents the chemical reaction leading to wax hardening or that physical factors are responsible. As noted previously, Beament suggested that a monolayer was largely responsible for prevention of water loss. If alcohols are absent from the wax, what molecules are present that would form a monolayer with

154

LAWRENCE I . GILBERT

TABLE XIV Composition of the cuticular wax of Periplaneta americana Chemical Class

% of wax

Components of class

Hydrocarbons

75-77

C27 n-alkene (heptacosa-9, 18diene) C25, C26, C27 n-alkanes C17-C20, C28, C29 n-alkanes C14, C16, C18 n-alkanoic acids C13, C15, C17 n-alkanoic acids C15, C16, C17 branched alkanoic acids C18 n-alkenoic acid (octadec-9enoic acid) unknown unsaturated C14, C15, C16 n-alkanals C13, C17-C25 n-alkanals C16, C18 branched alkanals C18 n-alkenal (octadec-9-enal)

Fatty acids

Aldehydes

Esters Sterols

7-1 1

8-9

Relative abundance of components weight % of class 66 15, 14, 12 each < 1 5 , 24, 23 each ,1 each 1-2 35 6 16, 10, 14 each 2-8 0.5, 3 3

3-5 <1

Unaccounted for 2-3 From Gilby and Cox (1963).

low permeability to water? Gilby and Cox argue that fatty acid monolayers would be ineffective in controlling water evaporation and suggest that either some unusual and uninvestigated property of the wax causes the formation of these hypothetical monolayers or that a three dimensional network exists in the wax which imparts water impermeability characteristics to the cuticle. This latter alternative may be due to associations of the aldehydic components of the wax, but both of these suggestions are without experimental proof. At least one insect does contain a large amount of alcohol (albeit solid) exterior to its cuticle. For those of us who have worked with the larvae of S. cynthiu, the nature of the crystalline white powder on the cuticle has proved somewhat of a mystery. Very recently, Bowers and Thompson (1965) have demonstrated that this material is composed

LIPID METABOLISM A N D F U N C T I O N I N INSECTS

155

of saturated alcohols. The major component is a C30alcohol (n-triacontanol) with traces of a C28alcohol (n-octacosanol). The triacontanol actually comprises about 93% of the total cuticular lipid. This finding is in sharp contrast to the composition of the cuticular lipids of the cockroach and points out how difficult it is to generalize in this research area. Injection of [l4C]acetateinto the Cynthia larva resulted in a small but significant amount of label in these alcohols suggesting that the larvae synthesize these long chain saturated lipids rather than the alcohols being a product of leaf ingestion. If these alcohols are secreted by the epidermal cells, then it is possible that such secretions may be under endocrine control and may find parallels in the work of Locke (1964) who has described a most intriguing phenomenon in larvae of Calopedes ethlius. After the moult to the fifth instar, the larva secretes wax in the form of cylinders, 0.5 p in diameter, through two patches of modified integument. This process seems to be under endocrine control. Although much important information has been gleaned regarding the morphology of the secretion and cellular elements responsible (cf. Locke, 1964), little or no biochemical work has been conducted on either the nature of the wax or its biosynthesis. If indeed the process of wax secretion is under endocrine influence, the biosynthesis of the wax may be an ideal subject for the study of hormones on biochemical processes. B . EXTRA-CUTICULAR

Hydrocarbons do exist in tissues of the insect other than those associated with the cuticle. For example, the scent secretion of the hemiptperan Rhoecocoris sulieventris contains n-tridecane as the major hydrocarbon component along with n-dodecane and n-undecane (Waterhouse and Gilby, 1964), and Baker et al. (1963) showed that the hemolymph of the Mormon cricket contains saturated hydroCzs with traces of C41to carbons with chain lengths of C25,CZ6,C27, C43. Acree et ul. (1965) have reported on their gas chromatographic analysis of the principal hydrocarbonsin the hemolymph of P. americuna and B. germanica. As an example of the almost unmatched flexibility of the gas chromatograph, these investigators were able to analyse the supernatant of centrifuged hemolymph by injecting it directly into the instrument. The three main hydrocarbons of male Periplaneta were pentacosane, methyl pentacosane, and heptacosadiene ; the latter constituting more than 50% of the total at all stages studied. This finding agrees with that of Baker et al. (1963) as does the observation that

156

LAWRENCE I . GILBERT

twenty-four-day old adults show a sexual dimorphism in hydrocarbon content. That is, the female contains more than the male, although the same relative concentration exists in both sexes of younger adults. It would have been of interest to determine whether this sexual difference also existed in castrated animals. The older females are presumably undergoing oogenesis and it is possible that this involves a decrease in the water content of the hemolymph. Since these investigators analysed a constant quantity of hemolymph without determining the blood volume, the possibility exists that the rise in the quantity of hydrocarbons per volume of hemolymph is an indirect result of “ desiccation” rather than a direct result of hydrocarbon synthesis. Perhaps the total quantity of hydrocarbon in the hemolymph remains the same in both sexes of older cockroaches, but greater loss of water occurs in the female. This problem of water loss assumes great importance in experiments dealing with hemolymph whether it concerns lipid, protein or any other chemical constituent. Many “alterations ” in concentration may be more apparent than real. The function of these hydrocarbons in the hemolymph is not known with certainty but they are most likely involved in the synthesis of new cuticular lipid after each moult. A novel problem relating to the biosynthesis of lipids by insects is that regarding the wax secreted by the honey bee (Apis rnelliferu) which is used for housing the brood and the storage of nutrient (honey). As Young (1963) points out, it is the young worker bees who secrete the major portion of the colony’s wax. By feeding bees heavy water and [1-l4C]acetate, Piek (1961) found that the acetate is incorporated into wax acids and hydrocarbons but not into the esters. He suggested that the FFA were synthesized from acetate in a tissue distinct from that whose function was the synthesis of esters and alcohols. Piek then postulates that the hydrocarbon portion of the wax could then be formed from the FFA by oxidation to ketones and subsequent reduction. By autoradiographic analysis, Piek (1964) demonstrated that the oenocytes take up labelled acetate whereas the fat body cells fail to do so. This is rather unusual in itself since acetate is readily taken up by the fat body of many other species of insects during fatty acid biosynthesis (see Section IV). Piek‘s supposition is that the oenocytes synthesize the wax acids and hydrocarbons from acetate derived from the fat body. The fat body cells derive the acetate from the glycolytic pathway and are responsible for providing the acetate precursor to the oenocytes, as well as synthesizing the esters and their component acids and alcohols from acetate. The suggestion that the oenocytes participate in the production of wax is not new since Wigglesworth (1947, 1948) has

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presented evidence that the oenocytes are responsible for the production of cuticular lipids. Further work is needed to substantiate this hypothesis but it is nevertheless an attractive theory.

VI. ISOPRENOID COMPOUNDS A . N U T R I T I O N A L STUDIES

Isoprenoid compounds are of vital importance to the insect. These include the steroids and terpenes as well as other compounds. [Terpenes are compounds chemically related to the simple isoprene, C5H8 and are usually classified as monoterpenes (C,,) ; sesquiterpenes (C15) ; diterpenes (C2,); triterpenes (C,,) (Nicholas, 1963)l. Since in most living things (and insects appear to be one exception), sterols and terpenes utilize the same biosynthetic pathway, these important molecules will be discussed together. It is a vast topic for a general review and several aspects will be omitted. Because of the importance of some of these substances as insect hormones or mimics of insect hormones, certain endocrine aspects will be alluded to although the reader should consult Gilbert (1964) for a comprehensive survey of this topic. An excellent review on the utilization of sterols by insects is that of Clayton (1964). For general reviews of cholesterol and isoprenoid metabolism, see Cornforth (1959), Wright (1961), Popjiik (1963), Goodwin (1963), Garattini and Paoletti (1963), Talalay (1957), Staple (1963), Nicholas (1963), Bloch (1965). See Lederer (1964) for a review of the origin and function of some methyl groups in branched chain fatty acids, plant sterols and quinones. Unfortunately there is no work discussed in which insects have been utilized. Since carotenoids and insect pigments will not be discussed the reader is referred to the reviews of Olson (1964), Cromartie (1959) and Fuseau-Braesch (1963). The formulae for several of the more important steroids relating to the current topic and that will be alluded to in the present discussion, are given in Fig. 18. One of the basic tenets in insect biochemistry is the inability of insects to convert simple precursors to sterol. This was first demonstrated by Hobson (1935a, b), who showed that the fleshfly Lucilia sericata, required an exogenous source of sterols for normal growth. This observation has been corroborated in a multitude of nutritional experiments on various insect species (cf. Clayton, 1964). There is no rule as to when a sterol deficiency will manifest itself, but varies in different insects. In some cases the nutritional deficiency can be clearly seen shortly after hatching from the egg, in Gthers just prior to pupation, and in still others, only in the process of oogenesis (Robbins and

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L A W R E N C E I . GILBERT

Cholesterol

Cholestanol

< 7-Dehydrocholesterol

< A5-Cholestanone

A5-Cholestene

7-H ydroxycholesterol

,k?-Sitosterol

Ergosterol

FIG.18. The structure of some sterols important to insects.

L I P I D METABOLISM A N D F U N C T I O N I N INSECTS

159

OH

0 Zymosterol

Ecdysone FIG.18 continued.

Shortino, 1962). In some instances, sterols are requisite for normal embryonic development and hatching (Chauvin, 1949;Monroe, 1959,1960). To explain this sterol requirement in the diet of insects, experiments over the past two decades have fallen into four main categories. First, a nutritional survey of many sterols and sterol precursors that can replace or “spare” cholesterol, the main insect sterol. Second, biochemical investigations aimed at proving that insects can or cannot synthesize sterols from simple precursors. Third, experiments on how insects (particularly phytophagous insects) can structurally modify the dietary sterols. Finally, more recent investigations have attempted to define some physiological role for these sterols. Noland (1954) found that various derivatives of cholesterol can be utilized by B. germanica. Nutritional studies on flies (Silverman and Levinson, 1954; Levinson and Bergmann, 1957), Dermestes larvae (Fraenkel et d.,1941, Clark and Bloch, 1959a, b, c), Cdobruchus larvae (Ishii, 1955) and a host of other insects (Table XV) conclusively show that a whole range of sterols can substitute for, or partially spare, cholesterol in the diet. In a classic paper on sterol utilization in Dermestes uulpinis, Clayton and Bloch (1963) demonstrated that cholestanol had a full cholesterol sparing activity. They postulate that any sterol that can fully replace cholesterol in the diet must be closely allied to cholesterol in structure so that it can fit into the correct “spaces”. The substituting molecule must be 3-/?-hydroxy and must have essentially the planar structure of cholesterol itself. The introduction of a A5 bond restores some of the activity lost as a result of the removal of successive carbon atoms from the side chain. The requirements for cholesterol sparing activity in insects as suggested by Clayton and Bloch are: (1) The molecule must have a generally planar nuclear stfucture. (2) It must have a side chain of the cholestane type. (3) It must possess a /? oriented 3-hydroxyl group. 7.

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LAWRENCE I . GILBERT

It appears that some insects can at least preferentially extract from their medium that sterol most closely resembling cholesterol. Even if several A5,3 8-hydroxy sterols are present in the defined medium of the housefly, it will selectively take up or retain that sterol most closely approximating cholesterol (Thompson et al., 1963). In a detailed investigation of the nutrition of L. rnigratoria and S. gregaria, Dadd (1960a, b) has shown that cholesterol will satisfy the sterol requirement of these insects. These orthopterans can also utilize cholesteryl acetate, /I-sitosterol and dihydrocholesterol but not stigmasteryl acetate, ergosterol, 7-dehydrocholesterol, cholestenone or 7-oxocholesteryl acetate. Dadd also concludes that an hydroxyl group at C3 that can be esterified is required but that a ketone group at C, prevents sterol utilization (see also Levinson, 1955, 1960, 1962). The ability of insects to live on sterol free diets has been reported, but without exception has been accounted for by the presence of digestive tract symbionts that do have the capacity to convert acetate to sterol (cf. Pant and Fraenkel, 1954). While on the subject of nutrition, it should be noted that in addition to their role as dietary factors, certain terpenoid derivatives including sterols appear to be important as phagostimulants (cf. Thorsteinson, 1960). A great deal of this work has utilized the commercial silkworm due to the persistent efforts of the Japanese in their search for a synthetic substitute for mulberry leaves. fi-Sitosterol has been implicated as both a phagostimulant and “biting factor ” (Hamamura, 1959; Hamamura et al., 1961; Hamamura and Naito, 1961; Ito et al., 1964). Hamamura believes that separate chemicals attract the silkworm larva to the food, cause them to bite it, and finally induce swallowing. Several of these may be terpenes in accordance with Watanabe’s (1958) data indicating that hexenol and hexenal act as attractants. In addition to j3-sitosterol, stigmasterol and campesterol may also act as phagostimulants (It0 et al., 1964). However, in grasshoppers it appears that the phagostimulatory action of wheat germ oil is a consequence of the PL present (Thorsteinson and Nayer, 1963). Waldbauer and Fraenkel(l961) are of the opinion that plant odours will cause insect larvae to bite on any leaf. Dadd (1963) suggests that biting appears to follow orientation to olfactory stimuli common to many plants and that it is an automatic process. This may be true since even Bornbyx larvae will feed on plants other than mulberry (Legay, 1958). Though the basic questions remain open, the role of isoprenoids as phagostimulants presents a most intriguing and challenging problem. It now appears that certain plant monoterpenoids may be in part

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161

responsible for sexual maturation of the desert locust (S. gregaria). A low titre of these essential substances is present in senescent leaves and may explain why sexual maturation is delayed in those locusts feeding upon older leaves (Ellis et al., 1965). B . ISOPRENOID BIOSYNTHESIS

When labelled acetate is administered to a vertebrate or incubated with vertebrate tissues, isotopically labelled cholesterol can usually be extracted from the incubation mixture. In higher animals we now know that every carbon atom in cholesterol is derived from either the methyl or carboxyl carbon of acetic acid (cf. PopjAk, 1963). The reason that insects require a sterol in their diet is they are unable to perform this same synthesis, owing to one or more metabolic blocks between acetate and cholesterol (Clark and Bloch, 1959a, b, c; Kodicek and Levinson, 1960; Robbins, et al., 1960; Schaeffer et al., 1965; Goodfellow and Gilbert, 1965). This failure to use acetate as a building block in cholesterol synthesis is not restricted to the Insecta but exists in crustaceans, araneideans and diplopods (Zandee, 1962, 1964), as well as other invertebrates (Wootton and Wright, 1960). Precursors other than acetate have been unsuccessfully tried in the hope that the metabolic block exists prior to the synthesis of the precursor. For example, farnesol could not substitute for cholesterol in the diet of the rice stem borer (Ishii and Hirano, 1961) and [14C]mevalonatewas not converted to sterol by the housefly (Kaplanis et al., 1961), Cecropia silkmoth (Goodfellow and Gilbert, 1965) and several other insects (cf. Clayton, 1964). Sedee (1961) has demonstrated that in aseptically reared Calliphora, squalene cannot replace cholesterol as a growth factor and that in the presence of labelled acetate, the cholesterol of the insect remains devoid of isotope. Figure 19 shows the biosynthetic scheme leading to cholesterol as we now understand it for higher animals and micro-organisms. This scheme is important because it shows the close relationship between sterols and non-sterolic isoprenoids as well as demonstrating a series of reactions that the insect is incapable of conducting. The first step in sterol biogenesis is the activation of acetate and subsequent condensation of acetyl CoA and acetoacetyl CoA to form 3-hydroxy-3-methylglutaryl CoA. This five carbon compound is then reduced by NADPH to form mevalonic acid. Mevalonic acid in the presence of mevalonic kinase and ATP yields mevalonic acid pyrophosphate. Decarboxylation and dehydration of this compound gives rise to isopentenyl pyrophosphate

c

Q\ h)

TABLE XV Sterols as growth factors in various species of insects'

Species

7DehydroCholes- choles- Choles- Ergosterol terol tan01 tan01

Lucilia sericata Dermestes vulpinus

-

++

-

+

0

0 0

0 0

0 0

++ ++ ++ ++ ++ + ++f +++ 0 +++ ++ ++f ++

+++

+ + ++ + 0

+++ +++ ++ +++ +++ ++

-

-

-

-

++f ++f +++ +++ 0 Tribolium confusum + + + + + + Lasioderma sericorne + + -k + + + Silvanus surinamensis + + f + + + Ptinus tectus + + + + + + Stegobium peniceum ++f +++ Ephesiia kuhniella Attagenus piceus

EpiCholest- CholestSito- Stigma- Stigma- choles- Copros- Zymo- 4-en4-en- Ergosterol sterol Stan01 tan01 tan01 sterol 3-one 38-01 sterol

Phormia regina ++f Musca vicina + + f Locusta migratoria

+++

0

+++

+++ ++ +++ +++

0

0

+

+++ ++

0

(0)

-

++ +

0

+++

+++

-

0

._

Schistocerca gregaria + Blatella germanica Gryllus domesticus Bombyx mori Aedes egypti Tenebrio molitor -t Cdosobruchus chinensis Pyrausta nubilalis Drosophila melanogaster Hylotrupes baJulus ++

++ +++ +++ +++ +++ + ++ ++ +++ ++

+++

+++ +

-

++ 0

+++ 0 ++ ++ ++ + +++

++ +-+

-

+++ +++ +++ +++ +++ +++

+++ +++ +

++

+++

-

-

-

-

0 -

tt+

++

0

+(0) 0

++ +++ 0 + +++ -

+++

+++ +

The symbols indicate good ( +), moderate (+ +), poor (+), or no (0) utilization of a sterol by a given species. Some negative results obtained with the sterol acetates are placed in parenthesis. (-) indicates “not studied”. (From review by Clayton, (1964); See Clayton’s article for original reference.)

I 64

LAWRENCE I . GILBERT

which is converted to its isomer y,ydimethylallyl pyrophosphate. [Isopentenyl pyrophosphate, sometimes known as "active isoprene" is not only a key intermediate in the biosynthesis of many terpenoids, but also in the synthesis of carotenoids (Goodwin, 1963). Since some insects appear unable to synthesize the requisite carotenoids for normal visual reception (Goldsmith et al., 1964), the metabolic block is most likely prior to isopentenyl pyrophosphate.] Condensation of two of these 5-carbon molecules yields geranyl pyrophosphate which, together with

C~ASH 0

OH

II

0

I

II

HOC-CH2C-CH2-C-SCoA 2 NADPH

OH

+H+

2 NADP

0

II

CH"C-0-

I

I1

xATP

2-03POCH2CH,C-CH,C-OAH, OH

JO~ADP II

I I

'-O8P20CH2CH2C- CH2C- 0CH3 s G 3-OJ'@CH2CHzC=CH,

I

~

+

C

O

~

. 'A

CH3C=CHCH20P,0,3C H S HP@?- CH, I CH,C=CHCH,CH,C= CH, I

CHCH,OP,0,3~ = ~ H , C H , O P z O ~ -

kH3

HPzO?-

H3

CH3-C=CHCH2CH2C=CHCH2CH,C=CHCH20P20~-

I

CH,

I

CH,

I

CH,

L I P I D METABOLISM A N D F U N C T I O N I N IN SECTS

CH,

2 HP,O,S-

165

CH3

CH, CH,

FIG.19. The biosynthesis of cholesterol. a. Acetoacetate + acetyl CoA combine to yield hydroxymethylglutarate in the presence of a condensing enzyme. b. Hydroxymethylglutarate NADPH yields mevalonate. c. Mevalonate ATP yields 5-phosphomevalonate in the presence of mevalonic kinase. d. 5-Phosphomevalonate + ATP yields 5-pyrophosphomevalonate in the presence of phosphomevalonic kinase. e. 5-Pyrophosphornevalonate ATP yields isopentenyl pyrophosphate COZ. f. Isopentenyl pyrophosphate + dirnethylallyl pyrophosphate combine to form geranyl pyrophosphate. Dimethylallyl pyrophosphate is an isomerization product of isopentenyl pyrophosphate. g. Geranyl pyrophosphate + isopentenyl pyrophosphate yield farnesyl pyrophosphate. h. Reductive dimerization end to end of two molecules of farnesyl pyrophosphate yields squalene in the presence of NADPH. i. Cyclization of squalene to yield lanosterol in the presence of oxygen. j. A series of reactions whereby lanosterol is converted to cholesterol via 1Cdesmethyllanosterol, zyrnosterol and desmosterol. These latter compounds have been omitted from the scheme for the sake of simplicity.

+

+

+

+

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LAWRENCE I . GILBERT

another molecule of dimethylallyl pyrophosphate, gives rise to farnesyl pyrophosphate. The condensation reaction to form farnesyl pyrophosphate is mediated by a farnesyl synthetase extracted from liver and yeast (Lynen et al., 1958). Condensation of two of these C15molecules leads to the formation of the dihydrotriterpene squalene (C3,). Folding of this molecule yields lanosterol and finally cholesterol. Along the pathway, dephosphorylation may yield such endocrinologically active molecules as farnesol (see Section VI). Before continuing our discussion of the sterols, let us briefly discuss another most important molecule containing isoprene units. This is coenzyme Q (CoQ) which plays a crucial role in electron transport and shares part of the biosynthetic pathway leading to sterol. The coenzymes Q (CoQ, to CoQ,,) differ from one another only in the length of the isoprenoid side chain, the quinone portion of the molecule being identical. Gloor and Wiss (1958, 1959) have shown that mevalonate is a precursor of the isoprenoid chain but is not involved in the synthesis of the quinoid nucleus. The entire isoprenoid side chain appears to be synthesized by a stepwise incorporation of C5 units derived from mevalonate. The mechanism by which mevalonate is converted to the C5 unit is now understood (Lynen, 1959). Mevalonate is decarboxylated and diphosphorylated to isopentenyl pyrophosphate. This molecule then isomerizes to form 3,3-dimethylallyl pyrophosphate which is subsequently alkylated by another molecule of active isoprene, yielding allylic pyrophosphate. Further alkylations by active isoprene increase the chain by one unit at a time. The final reaction is thought to be an electrophilic aromatic substitution on an unknown precursor by the isoprenoid allylic pyrophosphate (cf. Wagner and Folkers, 1964).

0 0

CH,O CH30

f"'

CH,

CH,CH=CCH,

) H

0 Coenzyme Q

That CoQ is an important constituent of the mitochondria is seen from the fact that its concentration in this organelle is higher than any of the cytochromes. Although Laidman and Morton (1962) demonstrated the presence of CoQB,CoQ9 and CoQ,, in larvae of the blowfly and suggest that they may have been products of digestion (see also Lester and Crane,

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167

1959; Heller et al., 1960), almost nothing is known regarding the possibility of CoQ biogenesis in insects. One can assume that insects most likely cannot synthesize this molecule if a metabolic block exists prior to active isoprene. We do not really know whether CoQ is as crucial to the proper functioning of the insect mitochondrion as it is to beef heart mitochondria, although we must assume for the present that it is. It would appear that an almost ideal situation for the study of both function and synthesis of CoQ in insects, would be tissues such as developing flight muscle where the giant mitochondria are undergoing morphogenesis. This would be especially true in some Lepidoptera where this morphogenesis occurs in a “closed system” and one need not worry about ingestion of materials containing exogenous CoQ. To return to the main topic of sterol biosynthesis, we have stated that insects cannot synthesize sterols. However, in the past several years there have been reports that either conclude or suggest that certain insects can convert acetate to sterol. Most of these data have been shown to be due to symbiotic micro-organisms and not to the insect’s ability. Because of the importance of this point we will discuss a few recent examples. When the silverfish Ctenolepisma sp. was reared on a diet containing [‘*C]acetate, some radioactivity was recovered in the sterol fraction (Clayton et al., 1962). These investigators did suggest that any sterol biosynthesis may be due to symbionts in this notoriously infected insect. When a closely related insect, Thermobia domestica, was grown aseptically and fed or injected with [l*C]acetate, only about 0.001% of the label was recovered in cholesterol (Kaplanis et al., 1963a, b). These authors doubt that the firebrat has the capacity to synthesize sterols from acetate and we must agree. When 1 to 2 pc of labelled acetate is injected into silkworm larvae or pupae, some radioactivity is recovered in the sterol fraction (Saito et al., 1963a). However, this is extremely low and ranges from 1-69-56.88 cts/min per mg of the digitonide. Although theseauthors argue that acetate is a sterol precursor in Bombyx, the percentage of the initial isotope recovered was extremely low. It was, in fact, of the same order of magnitude as that obtained by Sridhara and Bhat (1965b) in the same insect injected with labelled mevalonate, but they conclude “that the sterol biosynthetic pathway is absent in this insect”. However, even this extremely low quantity of incorporation could be evidence of synthesis. The major objection is that even this is most likely to be due to symbiotic micro-organisms. We have also investigated this problem of sterol biosynthesis in another silkworm, H. cecropia (Goodfellow and Gilbert, 1965). An

168

LAWRENCE I . GILBERT

aqueous solution of ~ ~- [ 2 - ~ ~C] me v a l onafe was injected into pupae and developing adults and the lipids analysed after adult emergence. About 8% of the label was incorporated into lipid and 93% of this was recovered in the non-saponifiable fraction. Column chromatography yielded a sterol fraction that was precipitated with digitonin. The digitonide was cleaved and the cholesterol purified. Only 0.011% of the radioactivity was associated with cholesterol. This low level of activity and our inability to obtain constant specific activity during the purification process suggests that there is no significant incorporation of mevalonate into cholesterol in this insect. C. I S O P R E N O I D C O N T E N T

It is almost 30 years since Bergmann (1934) reported that chrysalis oil (lipid extract of B. mori pupae) contains sterols composed of 85% cholesterol and 15% sitosterol (as we shall see, other sterols exist in this insect in addition to these two). Since that time, numerous insects and insect tissues have been analysed. Prior to the use of gas chromatography, large amounts of sterol were necessary for analysis. For instance, in their work on the sterols of M . domestica, Agarwal et al., 1961) extracted 65,000 houseflies and obtained 319 mg of crystalline sterols. Today, analyses can be accomplished on the tissue of a single insect (Goodfellow and Gilbert, 1965). Numerous isoprenoids have been identified in insect tissues. Some of these have their.source in the insect’s food, some result from the insect’s capacity to modify dietary sterols, while others are the result of the metabolic processes of the symbionts. The more common sterols will be discussed in detail subsequently but we may mention some of the more recent findings now. The major sterol of pupal and adult Colorado potato beetles is the common phytosterol p-sitosterol (Schreiber et al., 1961). Mass spectrometric analysis of the sterols of Calotermesflavicollis revealed the major body sterol to be cholesterol with lesser amounts of stigmasterol and p-sitosterol plus an as yet unidentified sterol (Duperon et al., 1964). Queen bee larvae, as well as the royal jelly upon which they feed, contain 24-methylene cholesterol (Barbier and Bogdanowsky, 1961; Pain et al., 1962; Weaver et al., 1964). It is of interest that this rather exotic sterol was also identified in the pollen collected by the bees, suggesting that the insect may have little to do with its formation. Wlodawer and Wisniewoska (1965) have identified a “fast acting” sterol in the hemolymph of Galleria larvae that may be 7-dehydrocholesterol.

L I P I D METABOLISM A N D F U N C T I O N I N I N S E C T S

169

The fact that some insects have come to depend solely on their natural food source for a specific sterol is demonstrated by Drosophila pachea which breeds only in the stems of the senita cactus. Heed and Kirscher (1965) could not rear this fly on a normal laboratory diet unless it was supplemented with cactus. Further investigation demonstrated that the natural sterol of the cactus, A7-stigmasten-3/3-ol (schottenol) was required for growth and although it could be replaced by A7-cholesten-3/3-ol or A5*7-stigmastadien-3-/3-ol;other sterols including cholesterol did not support larval growth. Nutritional studies indicate that this insect cannot demethylate at C24and differs from some other insects discussed previously. The above observation points out rather profoundly that few definite conclusions can be applied to metabolic processes in all insects. As workers investigate more insects with even more exotic ecological niches, further metabolic specializations will undoubtedly be discovered. The volatile terpenes of the termite soldier cast (Nasuititermes) have been studied by Moore (1964). The major component appears to be a-pinene in the defensive secretion of three species. B-Pinene and other monoterpenoid hydrocarbons are also present and may function as solvent carriers and perhaps alarm substances. Moore is of the opinion that the soldiers synthesize these compounds rather than obtain them through ingestion, but this has not been conclusively demonstrated. The mandibular glands of the workers of the formicine ant Acanthomyops claviger secrete a mixture of the stereoisomers citronella1 and citral in the ratio of 9:l (Chadha r?t al., 1962; see also Butenandt et a/., 1959a). Recently, Law et al. (1965) showed that ants of different sexes and castes produce different odorous compounds, the volatile components of which are composed of several simple terpenes or terpene derivatives. These mixtures are most likely used as mating pheromones and although the terpenes are qualitatively similar, each species of ant appears to produce a distinct blend. Among the compounds thus far In 1956, identified are citronellol and 2,6-dimethyl-5-hempen-l-ol. Kullenberg suggested that geraniol was present in the scent given off by the honey bee A . mellifera. By gas chromatographic analysis, Boch and Shearer (1962, 1963) showed the presence of geraniol in the Nassanoff organ of the bee. Their data indicate that the organ contains from zero to 1.5 pg of geraniol per bee. The fact that none is detected in newly emerged adults but is present subsequently, suggests that geraniol production is related to the physiological age of the bee. Whether any of these terpenes are in fact synthesized by the irfsect is a matter of conjecture.

170

L A W R E N C E I. GILBERT D . STEROL MODIFICATION

That a host of mechanisms exist for the metabolism of steroids is an established fact in mammals and micro-organisms. The fact that sixty steroids have been isolated from human urine alone attests to this statement (Talalay, 1957). In higher animals and micro-organisms the steroid skeleton can be oxidized, reduced or hydroxylated at several positions. In addition, they can be converted to lactones (Talalay et al., 1963). It would not be surprising if a great many more steroids exist in insects other than those already reported, since there are a great many ways in which steroids can be modified. According to Talalay (1957; Talalay et al., 1963) the principal metabolic means by which steroids can be transformed are : Interconversion of hydroxy and ketosteroids involving both the side chain and steroid rings. This necessitates a pyridine nucleotide-linked oxidation mediated by a hydroxysteroid dehydrogenase to give an unsaturated ketosteroid. There would then be an irreversible shift of the double bond to yield an a,,!?unsaturated ketone.

\

\

-C-OH-

-

C=O

(2) Introduction of a carbon to carbon double bond. -CHZ-CHZ-

-CH=CH-

(3) Hydroxylation of first, second or third degree carbons on either the steroid skeleton or side chain. \

CHz-

/

\ CHOH /

(4) Epoxidation of the ring of unsaturated steroids.

-CH=CH-

-

0

/ \

-CH-CH

(5) An oxidative break of carbon to carbon bonds resulting in ketones, acids and lactones. (6) Hydrolysis of steroid esters. (7) Conjugation of steroid alcohols to form sulphates and glucuronides.

LIPID METABOLISM AND FUNCTION IN INSECTS

171

Unfortunately, little is known regarding these possible means of steroid modification in insects but it is surely a fertile field for both the steroid chemist and insect physiologist. However, we do have convincing evidence that although insects cannot synthesize sterols, they can indeed modify them. For the most part, investigators of this phenomenon have used cockroaches, flies and moths, and only a few species of these. It would be presumptuous to extrapolate these data to the whole class. Agarwal and Casida (1960) first reported that cockroach nymphs can convert cholesterol to some unknown more polar sterol. By a variety of physical and chemical techniques, Vandenheuvel et al. (1962) showed that at least 99% of the sterol of Periplaneta appears to be cholesterol, whether the cholesterol is fed or injected. This confirms similar observations on the housefly (Kaplanis et al., 1960) and German cockroach (Robbins et al., 1961). In Periplaneta, almost all the cholesterol is unesterified (vroman et al., 1964). This is of some interest since it is most likely that cockroach nymphs can convert cholesterol into a more polar sterol but probably only a minute amount is involved in this transformation. In order for the nymphs to moult they must secrete a critical titre of ecdysone (see Section VI), which is a very polar steroid. If cholesterol is the only sterol source in the diet and the animal is incapable of synthesizing sterol from simpler molecules, it follows that cholesterol must be the precursor for the more polar ecdysone. In contrast to Musca, B. germanica apparently has the capacity to convert /?-sitosterol to cholestero!. When tritiated /?-sitosterol was fed to these cockroaches for 42 days, 80% of the label was recovered in cholesterol (Robbins et al., 1962). It thus appears that Blattella can remove the 24-ethyl group from the side chain of /?-sitosterol. In 1963, Clayton and Edwards utilizing 3H and I4C-labelledsterols, investigated the metabolic conversion of cholestanol to A'-cholestanol in aseptically reared B. germanica. They reported that this conversion involved the loss of 7/? and 8s hydrogzn atoms and suggested that the A7 sterol may possibly be formed in the digestive tract and distributed to other tissues from this synthetic site. They were, however, unable to identify 7dehydrocholesterol as a metabolite of cholesterol. 7-Dehydrocholesterol had been considered the major metabolite of cholesterol in insects. The work that led to this conclusion in almost all cases utilized insects raised under non-aseptic conditions, and could possibly be attributed to contaminating micro-organisms. However, Robbins et a/. (1964) reared Blattella nymphs under aseptic conditions on a semi-defined medium containing [4-14C]cholesterol and extracted the insects one to two

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months later. By infra-red, ultra-violet and gas chromatographic analysis, they showed that this roach did indeed convert the labelled cholesterol to labelled 7-dehydrocholesterol but not to ergosterol or 22-dehydrocholesterol. These latter two sterols were identified by Clayton (1960) in Bluttellu grown under non-aseptic conditions. Whether grown aseptically or not, about 3% of the total sterol of Bluttellu is 7-dehydrocholesterol. Thus, we can conclude that the tissues of the roach are capable of this conversion but that micro-organisms are most likely responsible for the transformation of cholesterol to ergosterol and 22-dehydrocholesterol. The significance of 7-dehydrocholesterol is unknown although Robbins and his colleagues (1964) suggest that it may act as a precursor for vitamin D or a similar compound. These investigators raise the important point that 7-dehydrocholesterol may be an intermediate in the conversion of cholesterol to ecdysone since 25% of the sterol in the prothoracic glands of P . umericanu during the terminal nymphal moult is 7-dehydrocholesterol. Further work on this subject is eagerly awaited. Kodicek and Levinson (1960) have suggested that blowfly larvae can subsist on j3-sitosterol as the sole sterol in the diet and postulate that this insect has the ability to de-ethylate the /3-sitosterol side chain at CZ4just as Bluttellu can demethylate ergosterol (Clark and Bloch, 1959a, b, c; Clayton, 1960). By the use of 3H-labelled /3-sitosterolY aseptic rearing conditions and chemical analysis, Kaplanis et al. (1965) showed that larvae of M . domesticu cannot dealkylate P-sitosterol to cholesterol. [The sterol of Muscu which was formally named muscasterol is actually a mixture of 74% campesterol and 21% /3-sitosterol (Thompson et ul., 1962)]. In addition, this sterol does not fulfil the sterol requirements of the fly. Adult female flies do have the capacity to dehydrogenate the /?-sitosterol to its corresponding 5,7 diene and although this is a major metabolite of A5 sterols in the adult female and egg, it is at most a minor pathway in larvae. During metamorphosis then, this sterol dehydrogenation pathway comes into its own and may be under hormonal control, although no work on this subject has been reported. Kaplanis and his co-workers suggest that previous reports indicating a sterol sparing action of /3-sitosterol in flies may be explained by any of the following: a biochemical difference in strains of flies; some contamination of the sitosterol by cholesterol (since only minute amounts of cholesterol are needed to fulfill the sterol requirement) ; presence of trace quantities of cholesterol in dietary ingredients such as casein; an efficient sterol storage and transfer mechanism by which the adult female can supply adequate amounts of cholesterol to

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larvae via the eggs. This investigation points out the difficulties of accepting that nutritional work where extraordinary precautions were not taken to ensure complete quality control. The principal sterols of Tribolium confusum larvae are cholesterol and 7-dehydrocholesterol. Recently, Smissman et al. (1964) have isolated and characterized dehydroepiandrosterone, pregnenolone and progesterone as the metabolic products of normal sterol metabolism in these insects. This indicates that Tribolium possesses the enzymatic capacity to alter the side chain of dietary sterols such as ergosterol which was in the yeast containing diet. These investigators tentatively identified androstenedione as well and proposed the following scheme for sterol interconversion in Tribolium : Ergosterol

Cholesterol

-

Dehydroepiandrosterone I I

+ I

Pregnenolone

I

Androstenedione + - - - - - - - - Progesterone

Whether any of these compounds that play such an important role in mammalian regulation have a physiological role in insects is an open question at this time. phytosteLevinson (1960, 1962) believes that the conversion of Czs-zs rols to cholesterol is probably a general characteristic of phytophagous insects, whether they are coleopterans, dipterans, hymenopterans, orthopterans or lepidopterans. This is accomplished by either the removal of the side chain of 6-sitosterol and de nouo synthesis of the cholesterol side chain or elimination of the supernumerary carbon atom to yield the iso-octyl side chain. On the other hand, obligatory carnivores are not required to have this ability since they receive cholesterol in their diet. More specifically, the conversion of phytosterols to cholesterol would involve the following molecular modifications according to Levinson : (1) Elimination of the methyl group of campesterol or the ethyl group of j3-sitosterol from G4.

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(2) Saturation of the A23:24double bond together with the elimination of the C24ethyl group of stigmasterol. (3) Saturation and de-ethylation of the side chain together with alteration of the position of the double bond in ring B of a-spina-sterol. Thus far, most of the evidence for these suggestions is circumstantial. That is, conversion does take place, but no experiments on the molecular level have been reported. In a series of experiments in which ergosterol was placed on leaves and fed to fourth instar Bombyx larvae, it seems that ergosterol is converted to cholesterol (Sridhara and Bhat, 1963). This is based on the indirect evidence that no ergosterol could be identified in the insect which itself showed an increase in the cholesterol content of various tissues. However, with this and almost all other work involving the Lepidoptera, one cannot discount the action of numerous symbiotic micro-organisms. The mulberry leaves upon which silkworm larvae feed contain p-sitosterol as the major sterol and lesser quantities of campesterol (Miyauchi et al., 1964). Bombyx contains cholesterol, fl-sitosterol and campesterol at all stages of development (Saito et al., 1963b). In no case however does campesterol exceed 7.8% of the total sterol. During the late larval instars, in pupae and in adults, the percentage of psitosterol decreases with a concomitant increase in cholesterol, indicating sterol interconversion. Of particular interest is their finding that the highest percentage of cholesterol exists in newly hatched larvae, indicating to the authors, a synthesis of cholesterol during embryonic development. Our studies (Goodfellow and Gilbert, 1963, 1965; Gilbert and Goodfellow, 1965) on H . cecropia also demonstrate the conversion of phytosterols and are similar in several respects to those just discussed on B. mori. We have found that cholesterol is the major sterol in all tissues investigated at all stages of development. Also present are the phytosterols, 15-sitosterol and campesterol and possibly minute quantities of stigmasterol as well as unidentified polar steroids (see Section VIF). The newly hatched first instar larva exhibits the highest relative concentration of cholesterol and we believe that this is due to selective uptake by the developing ovary from the female’s sterol supply. When the larvae begin to feed, the relative concentration of cholesterol decreases dramatically as the phytosterols predominate. Throughout metamorphosis, however, there is a progressive increase in the concentration of cholesterol relative to phytosterol until the five-day-old

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adult moth yields upwards of 85-90% cholesterol. The absolute concentration of sterol remains more or less constant during the pupaladult transformation. The fact that females have higher sterol titres than males suggests the necessity for sterols in oogenesis. The injection of [4-14C]cholesterolinto the hemocoele of developing adults resulted in the retention of unchanged cholesterol in various body tissues except for conversion of small quantities to polar steroids. Less than 2% of the cholesterol became esteriiied with fatty acids. Though we have repeatedly attempted to identify cholesterol in the food plant of Cecropia (Acer negundo), we have been unsuccessful although j3-sitosterol, campesterol and stigmasterol were identified. This demonstrates that H . cecropia does have the capacity to convert phytosterols to cholesterol (see also Schaeffer et al., 1965; Von Ardenne et al., 1965). That insects are capable of converting free sterols to sterol esters has been demonstrated repeatedly (Casida et al., 1957; Clement and Frisch, 1946; Kaplanis et al., 1960; Robbins et al., 1961;Clayton and Edwards, 1961). In a study of the sterols and sterol esters of female houseflies and eggs, it was found that sterol esters comprise 41% of the total sterol of the eggs but only 8.4% of the sterol of the adult female (Dutky et al., 1963; see also Kaplanis et al., 1960, 1963b). More than 90% of the fatty acid constituents of the sterol esters were c16 and C,, monounsaturated with the higher molecular weight fatty acid amounting to about 78% of the total. The fatty acid moiety of the egg and female TGL differed from those of the sterol esters in that the former were more saturated and fatty acid predominated. This is similar to the findings of Bade and Clayton (1963) who demonstrated that cholesteryl oleate is the major sterol ester of the cockroach E. floridana reared on a semisynthetic diet containing sodium oleate. Other sterol esters identified by these latter investigators contained linoleate and palmitate. In the eggs of Cecropia, about 36% of the cholesterol is esteriiied (Goodfellow and Gilbert, 1965). The function of these esters in the egg and developing embryo is unknown. We can conclude from the foregoing that insects require an exogenous supply of sterol for the fact that they are incapable of producing their own. Why is it vital for many insects to have sterols? What then are the functions of isoprenoids in living things? E . FUNCTION

The relatively constant concentration of sterols in insect tissues throughout development and the requirement for sterol in the diet

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suggest that the bulk of the dietary sterol is utilized as elements of cellular structure (cf. Clayton, 1964). There is good evidence that unesterified cholesterol plays an indispensable role in the maintenance of cell membranes in mammals (cf. Ball and Joel, 1962). Perhaps this is the primary role in insects as well. In support of this view, an analysis of the tissues of E. floridana by Clayton and Lasser (1964), and those of H. cecropia (Goodfellow and Gilbert, 1965) by differential centrifugation, demonstrated that the free cholesterol of insect tissues is almost entirely associated with particulate fractions. In both insects the free cholesterol pool has an extremely low turnover rate, also suggestive of a structural role for cholesterol. This is not, however, the only role for cholesterol in insects as it appears to be a precursor for at least one insect hormone (see Section VI F). We know very little as to why insects are unable to synthesize sterols. Either they do not possess the appropriate enzymes or contain substances that inhibit one or more processes in the biosynthetic pathway. Perhaps insects are capable of synthesizing geranyl pyrophosphate and farnesyl pyrophosphate (compounds that could be converted to farnesol and geraniol). This may result in the formation of terpenoid acids by oxidation of these terpenes (e.g. farnesol to farnesal to farnesoic acid). Reactions such as these have been demonstrated in the soluble fraction of mammalian liver (Popjiik et al., 1959). The accumulation of these terpenoid acids may exert some control over sterol biogenesis by inhibiting mevalonic kinase (Wright and Cleland, 1957; Levy and Popjiik, 1960) and result in accumulation of mevalonate. Thus, the accumulation of terpenoid acids may block sterol biosynthesis in insects. It has been shown that extracts of liver mitochondria have an inhibitory effect on cholesterol biosynthesis in rats both in uiuo and in uitro (Migicorsky, 1960, 1961, 1964). The inhibitory material is bound to a protein and it is postulated that this is one means by which cholesterol biosynthesis is controlled in the animal. Or, it could be that inhibitors such as this are present in insect tissues and prevent these animals that are so capable of synthesizing the most exotic chemicals from synthesizing the sterol nucleus. Unfortunately, no evidence is available to support either of the foregoing suggestions. F. INSECT HORMONES

Although this is not the proper place for a review of insect endocrinology (see reviews by Gilbert, 1964; Schneiderman and Gilbert, 1964), certain aspects should be discussed since they are intimately correlated

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with insect lipids. First, some of the insect hormones and pheromones are lipoidal. Secondly, at least one of these hormones affects the insect's ability to metabolize lipids. Finally, it is the author's belief that much of the future work in insect lipid chemistry and biochemistry will deal with the biosynthesis of these humoral factors as well as the mechanisms involved in catabolizing them to endocrinologically ineffective molecules. As is well known, moulting in insects is controlled by neurosecretory cells of the brain that release the brain (ecdysiotropic)hormone which in turn activates the prothoracic (ecdysial) glands. These latter glands then release the moulting hormone (ecdysone) which initiates the moulting process along with its manifold biochemical and genetic adjuncts. The corpora allata secrete the juvenile hormone, which, by favouring the synthesis of larval structures, prevents adult differentiation. In many adult insects the corpora allata regain their activity and release a gonadotropic hormone that is most likely identical with the juvenile hormone. The gonadotropic hormone is responsible for the completion of oogenesis in many adult females in addition to having other effects. Thus, these three endocrine glands are directly or indirectly responsible for many aspects of growth, differentiation and maturation. Let us now look at the current status of the chemistry of these hormones and any possible effects they may have on lipid metabolism. 1. Brain hormone

The purification. of the brain hormone has become a subject of confusion in the last few years. The first active material was an oily extract prepared by Kobayashi and Kirimura (1958) from 8,500 brains of B. mori. The hormone was soluble in organic solvents and appeared to be a lipid. Of interest were the reports by Kobayashi and his colleagues (Kirimura et al., 1962; Kobayashi et al., 1962) on the identification of the brain hormone. From 220,000 brains they prepared an oily extract which after several purification steps yielded 4 mg of crystals. The melting point of these crystals was 142"-143" and injection of 0.02 pg of this material dissolved in aqueous ethanol caused development in previously de-brained Bombyx pupae. From infra-red analysis and gas chromatography they concluded that the active material was cholesterol, or that cholesterol is a major constituent of the brain hormone. Positive assays were obtained with cholestanol, 7-dehydrocholesterol and in higher concentrations even 8-sitosterol, stigmasterol and ergosterol were effective. Once these results were published, numerous laboratories attempted

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to confirm them. In our laboratory, we have shown that in some cases as little as 0.005 pg of commercial cholesterol or sitosterol will cause development in debrained, diapausing saturniid pupae (cf. Schneiderman and Gilbert, 1964). The results, however, are erratic and no doseresponse curve could be obtained. Perhaps it is not the specific chemical structure of these sterols which is crucial for stimulation of the prothoracic glands, but physical factors such as surface tension or other properties of these solutions and suspensions. Our results on sterol identification in H. cecropia and several other saturniid moths, by micro-chemistry and gas chromatography strongly indicate that the brain hormone is not cholesterol (Goodfellow and Gilbert, 1963, 1965). These data reveal relatively high titres of sterol in the hemolymph of both male and female pupae (e.g. 587 pg sterol/ml hemolymph in female Cecropia), the major fraction of which is cholesterol. About 90% of the cholesterol is free rather than esterified, although it appears to be transported as a complex of cholesterol-hemolymphprotein. Thus, the amount of cholesterol normally present in the pupa’s hemolymph is 10,OOO times that necessary to stimulate the prothoracic glands. Although about 1% of the lipid extracted from the Cecropia nervous system is cholesterol (about 7% esterzed) it is for the most part associated with cellular and subcellular membranes and is most likely a structural component rather than a hormone (see also Clayton, 1964). In addition, the cholesterol of the insect nervous system has a relatively low rate of turnover (Goodfellow and Gilbert, 1965). That insects can convert phytosterols to cholesterol is no longer in doubt. The hypothesis that the brain hormone is cholesterol, is however very much in doubt. The medial neurosecretory cells of the brain of three species of mosquitos have recently been implicated in the regulation of aspects of lipid synthesis (Van Handel and Lea, 1965). Using whole insects, it was demonstrated that surgical removal of the neurosecretory cells within one hour after adult emergence increased the mosquito’s capacity for glycogen storage and decreased the normal tendency to store TGL. However, implantation of medial neurosecretory cells into previously operated animals does not restore metabolism to the normal level, while severing the medial neurosecretory cells from their axons has the same effect as cell extirpation. It is therefore possible that a neural rather than neuroendocrine explanation can account for the above results. Nevertheless, the differences between experimentals and controls was great and similar experiments on other insects will no doubt be conducted shortly.

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2. Moulting hormone (Ecdysone) (a) Structure. Of the three growth hormones known in insects, the purification of the moulting hormone has the longest history. There was more than a two decade span between the time that the initial chemical work began and the empirical formula was finally elucidated. Very recently, Karlson and his colleagues (1965; Karlson, 1963; Huber and Hoppe, 1965) reported the empirical formula of ecdysone as C2,H4406 with a molecular weight of 464. By a combination of chemical analyses and X-ray studies, it was established that this most important molecule was a steroid. Further analysis led to the structure of ecdysone (Fig. 18) as a very polar steroid containing 5 hydroxyl groups. (b) Synthesis. If ecdysone is a steroid, one may ask how it is synthesized by the insect. Since insects cannot synthesize steroids from more simple molecules, it is probable that cholesterol is a precursor of ecdysone. A preliminary experiment to test this hypothesis has been reported by Karlson and Hoffmeister (1963) who injected tritiated cholesterol into CaZZiphora larvae and extracted crude ecdysone. This material was radioactive and co-crystallizationwith unlabelled ecdysone yielded pure ecdysone of constant specific activity. Although the experiment is preliminary there is no doubt that cholesterol is a precursor of ecdysone in many insects. Since several species survive on a defined diet containing only cholesterol as the sterol nutrient, it follows that this sterol must be the building block for the synthesis of the moulting hormone. Our experiments on H. cecropia (Goodfellow and Gilbert, 1965) support this argument. When [4-14C]cholesterol or &~itosterol-~H is injected into the hemocoele of pupae, one always finds a small portion of the label associated with yet unidentified polar steroids (see also Kaplanis et al., 1960; Robbins et aZ., 1961). One of these may possibly be ecdysone and, although this has not yet been demonstrated, the fact remains that cholesterol can be a precursor of polar steroids, and ecdysone is extremely polar. It is of some interest that the highest concentration of these polar steroids was recovered in the intestine of H. cecropia. These compounds were neutral; that is, no bile acids or other acidic compounds could be detected, and no [14C]coprostanol, the major sterol excretory metabolite of mammals, could be demonstrated (see also Robbins et aZ., 1961). If some of these unidentified polar steroids are excretory metabolites, which seems likely, then the pathway of sterol degradation and thus ecdysone catabolism is quite different in this insect than in mammals. Elucidating the pathway of ecdysone synthesis and catabolism will no doubt "be actively pursued by biochemically oriented students of growth.

180 LAWRENCE I . GILBERT As regards the mode of action of ecdysone, it is now commonly believed that the primary effect may be on the genetic material (cf. Gilbert, 1964). However, all lipid changes during moulting and metamorphosis that were discussed previously are no doubt the indirect result of the moulting hormone. If, as Kroeger (1963)suggests, ecdysone acts upon the genes by influencing the sodium/potassium ratio in the nuclear sap of cells in the salivary glands of Chironomus, then ecdysone may exert its effect by acting on lipoidal components of the cell membrane or nuclear envelope. One may speculate that the sodium pump of which PL may be an integral component (see Section IV), is one target of the moulting hormone. This is a promising area of research with large dividends for the imaginative and patient investigator.

3. Juvenile hormone (a) Purzjication. The first active extract of juvenile hormone was prepared by Williams (1956)from the abdomens of adult male Cecropia moths. He showed that the active principle could be extracted from the tissues with ether and was neutral, heat-stable and oil-soluble. When this extract was injected into pupae, it ,duplicated in detail the effect of implanting corpora allata. That is, the pupae moulted into second pupae. To ascertain the nature of the active material, Schmialek (1961) extracted 80 kg of Tenebrio faeces with organic solvents and chromatographed the resulting oily material. From this and subsequent procedures he obtained 60 mg of oil with juvenile hormone activity. The active substances were identified as the terpene farnesol and its oxidation product farnesal. It appears that the biological activity of farnesol is dependent upon the trans configuration at the A6 or middle linkage (Yamamoto and Jacobson, 1962). Since that initial discovery, a large number of alcohols, alcohol methyl ethers and isoprenoid alcohols have been shown to possess both juvenile hormone and gonadotropic activity (cf. Bowers and Thompson, 1963; Schneiderman and Gilbert, 1964). [It is of interest that farnesol has also been implicated as a possible sex attractant of the male bumble bee (Stein, 1963)l. In 1963,Schmialek reported that [1-14C]mevalonate was incorporated into farnesol by S. Cynthia and intimated that farnesol or a derivative thereof is the juvenile hormone. Although this claim has been echoed by others, farnesol cannot be the juvenile hormone for the following reasons. We have demonstrated that the tissues of H. cecropia contain several compounds possessing juvenile hormone activity, including farnesol and phytol (Goodfellow and Gilbert, 1963, 1965). There are about 5 to 15

181 pg of farnesol (predominantly the trans-trans isomer) per gram of lipid in male moths and 20-25 pg of phytol (Fig. 20). This is not surprising when we consider that this insect is phytophagous and that farnesol is a common plant isoprene while phytol is a product of the disruption of the chlorophyll molecule. However, the same quantity of farnesol (if not more) is present in the chilled pupa, a stage containing no extractable juvenile hormone (Gilbert and Schneiderman, 1961b). Thus, approximately equal amounts of farnesol are present in the adult moth which stores large amounts of juvenile hormone and in the pupa which stores none. Although farnesol is present in the pupa, the pupal lipid shows no juvenile hormone activity because it takes milligram quantities of farnesol to give a positive bioassay whereas purified juvenile hormone is active in nanogram amounts. Secondly, we allatectomized male pupae and allowed them to complete adult development. Since the source of the juvenile hormone was removed, the lipid extracted from these moths was devoid of juvenile hormone activity, but contained the same quantity of farnesol as normal controls. The subsequent discussion of the purification of the juvenile hormone will add additional support for the postulate that farnesol is not the juvenile hormone, but only a mimic of the juvenile hormone. Notwithstanding the fact that the juvenile hormone may not be an isoprenoid, these compounds are of real interest. As stated previously, we do not know whether insects are able to synthesize isoprenes. When labelled mevalonate is injected into H . cecropia, less than 1% of the label is recovered in the phosphorylated isoprenol fraction (Goodfellow and Gilbert, 1965). Even this slight incorporation may be due to symbionts. What then is the current status of the juvenile hormone? By the time this review appears, we may know the chemical structure of this very potent molecule, but at this writing we do not. However, progress has been made since Williams’ original extraction. Interestingly enough, all the work thus far published on the purification of the juvenile hormone has used abdomens of the male Cecropia moth as starting material. Williams and Law (1 965) demonstrated a 4,000-fold concentration by repeated methanol extractions, saponification and silicic acid chromatography. By gas chromatographic techniques they recovered a fraction that was active at concentrations of 0.3 to 0.4 pg per 5 g assay pupa; a 50,000 fold purification. Mass spectrometric analysis revealed the major component to be an epoxide of methyl hexadecanoate, but pure isomers of this compound were devoid of juvenile hormone activity. LIPID METABOLISM A N D FUNCTION IN INSECTS

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Adult disoprenols

Diopouse disoprenols

e

f

FIG. 20. Gas-liquid chromatographic separation of the isoprenols of H. cecropia. Sample run on 10% ethylene glycol adipate column on washed silanized Chromosorb W (80/100 mesh). Column was 6 feet x 1/4 inch 0.d. Temperature 170°C and argon flow rate 60 ml/min, 20 psi. Gas chromatograph was an F and M with flame ionization detector. Lower graph represents isoprenols from a diapausing male pupa. a. trans-Nerolidol. b. cis-trans-Farnesol c. trans-trans-Farnesol d. t-Tetraprenbl ( G o ) cis-trans e. Not identified f. Not identified Upper graph represents isoprenols from a male moth. a. trans-trans-Farnesol b. t.-Tetraprenol (Ca0)cis-trans c. Not identilied d. Geranyl-linalool (C20) trans-trans (From Goodfellow and Gilbert, 1965.)

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These investigators suggest that the hormonally active molecule constitutes only a minute part of the most active fraction. However, their purified fraction was at least 1,000 times more active than any of the various isoprenoids reported to have juvenile hormone activity (see also Gilbert, 1964). Thus, almost a decade of work by this group has yielded a potent concentratebut not the structureofthejuvenile hormone. By the use of thin layer and gas chromatography as the principal tools, Roller et al. (1965) have also been able to prepare a concentrate from Cecropia oil that appears to possess about 100,OOO times the activity of the crude extract. Of interest is their finding that standards of farnesol, farnesal and farnesyl methyl ether all are eluted from the gas chromatograph much earlier than the true juvenile hormone peak, adding support to the argument that none of the above are true juvenile hormone. Our group (Meyer, Schneiderman and Gilbert) has been labouring with this same problem for almost a decade and can attest to the difficulties encountered. Among the problems are the difliculty of obtaining some 50,000 male Cecropia moths, the presence of mimicking substances in the original extract that may in some cases yield false positive bioassays, the inverse relationship between purity and stability, and finally the failure of the bioassay at inauspicious moments. Despite these obstacles, considerable progress has been made. Preliminary separations were conducted by absorption chromatography (alumina, zinc carbonate, silica gel). Up to a 75-fold enrichment was achieved by these procedures (Meyer and Ax, 1965a). It is of importance that extracts of five species of Lepidoptera showed similar elution properties, confirming in chemical terms what surgical experiments had suggested; namely that the juvenile hormone molecule may be the same in all insects (see however Slhma and Williams, 1965). In 1961, we reported a 100-fold concentration of the juvenile hormone by two successive counter-current runs of 50 transfers each (Gilbert and Schneiderman, 1961a). Using 300 transfers, Meyer and Ax (1965b) demonstrated a single band of juvenile hormone activity with a partition coefficient approaching unity when the solvent system was hexane : 93% ethanol, and 2.3 when the solvent system was petroleum ether: 83% ethanol. These results agreed with those of Gilbert and Schneiderman, (1961a), and yielded a very potent extract. Again, juvenile hormone extracts of H. cecropia and S. Cynthia ricini reacted similarly. In accordance with the now common practice, our most concentrated extracts were then analyzed by gas chromatography (Meyer et al., 1965). On the basis of these experiments and those alluded to above, it ~+A.I.P.

4

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LAWRENCE I . GILBERT

was concluded that our most potent samples had been enriched some 300,000-fold and were about 90% pure. On this basis it appears that a single male adult Cecropia abdomen contains, at most, 1-3 to 0.3 pg of juvenile hormone in its 200 mg of lipid. The minimum molecular weight of the molecule is about 300 and it is more polar than cholesterol. The injection of 3.3 nanograms of this concentrate into an assay pupa had the same effect as injection of 1 mg of crude extract. In our assay, farnesyl methyl ether, one of the more potent mimics of the juvenile hormone, shows a value of about 1,500 juvenile hormone units. The concentrated extract on the other hand gives a value of about 300,000,000 juvenile hormone units on the same scale. To summarize, one can say with some assurance that although the exact structure of the juvenile hormone is not known, it is surely none of the pure chemicals thus far described in the literature. (b) Mode of action. In our consideration of the mode of action of the juvenile hormone, we will discuss only those processes involving lipid metabolism although the main target of this hormone may be similar to that of ecdysone. By excluding all processes except those bearing on the main topic of this paper, we essentially restrict the discussion to that of the gonadotropic hormone. The basic observations made were that allatectomy of female adult flies, grasshoppers and cockroaches resulted in hypertrophied fat bodies, due mainly to an increase in lipid content (cf. Gilbert, 1964). Some investigators suggested that this was due to the failure of the hormone to cause utilization of the stored lipid in the fat body; others that the hormone acts directly on the ovaries and that fat body hypertrophy was an indirect consequence of the failure of complete oogenesis ; while still others suggested that the hormone secreted from the corpora allata had general metabolic effects. All of these observations and hypotheses are based on surgical experiments and lipid determination by gravimetric means. Is there evidence from higher animals that hormones do affect lipid metabolism? Vertebrate hormones most definitely affect lipid metabolism. The hypophysectomy of rats primarily affects the conversion of acetate to longer chain fatty acids either by a reduced capacity to take up acetate or convert it to acetyl CoA (Manchester, 1963). In rat adipose tissue, the lipase is sensitive to various hormones including ACTH, TSH, glucagon, etc. (Vaughan et al., 1964). Recently, Rudman (1963) has reviewed the adipokinetic action of hormones on mammalian adipose tissue. He defines adipokinetic as “the capacity of a substance to cause release of TGL in adipose tissue, with subsequent transport of the mobilized lipid by the blood to other organs” (p. 119). At least 10

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different substances isolated from the anterior or posterior pituitary, pancreas or sympathetic nervous system possess adipokinetic activity and all play important roles in mammalian homeostasis. As far as insects are concerned, we know that allatectomized P. americana females contain more lipid than control cockroaches (Bodenstein, 1955). This phenomenon has been recently investigated utilizing [l-14C]acetate(Vroman et al., 1965). They corroborated Bodenstein’s original observation in that allatectomized females yielded 66% more lipid than non-operated controls. Of the total lipid, the allatectomized insects contained 68% TGL while the controls had 52%. However, the allatectomized roaches contained 18% PL in contrast to 32% in the controls. On an absolute basis, the surgically deprived insects had twice the quantity of TGL as did the controls while the PL content was about equal. Vroman and his colleagues concluded that “the corpora allata regulate the metabolism of PL and TGL by controlling the mechanisms responsible for the utilization of lipids” (p. 903). Although this may be true, it is far from a proven fact. For one thing, allatectomy may have affected the animal’s feeding behavior and thus influenced the lipid content. It would be of interest to know whether implantation of active corpora allata into allatectomized animals brought the lipid picture to a normal level. Our results on the cockroach Leucophaea maderae (L. I. Gilbert, unpublished observations) support the conjecture that the corpora allata control aspects of lipid metabolism. In these in vitro studies, we found that active corpora allata (from H . cecropia or L. maderae) stimulate the incorporation of [l-14C]palmitiateinto ovarian glycerides as well as stimulate fatty acid oxidation. It is these neutral lipids that will subsequently provide some of the energy for embryogenesis. When isolated fat body is assayed in the same manner, we find that the corpora allata decrease the rate of biosynthesis of lipid and also the oxidation of fatty acids. The above results were confirmed with both juvenile hormone extract and farnesyl methyl ether, although their insolubility in water makes in vitro experiments somewhat difficult. The most consistent results were obtained with the very active corpora allata of newly emerged Cecropia moths. These in vitro studies that bypass the complexity of the whole insect, suggest that the corpora allata act on both the fat body and ovary of female Leucophaea by making more lipid available for storage in the maturing oocytes. This is not to say that the hormone has no effects on other metabolic processes, but its effect on lipid metabolism is certainly one reason the corpus allatum is an indispensable adjunct to oogenesis.

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4. Sex attractants

Since Butenandt and Tam (1957) isolated the acetate of trans-2hexen-1-01 from the abdominal gland of the tropical water bug, Kethocerus indicus, and showed that this material made the female more receptive to the male, the field of isolation and identification of lipoidal substances having pheromonic effects in insects, has progressed at a rapid rate. For recent reviews the reader is referred to Gilbert (1964), Jacobson (1963) and Jacobson and Beroza (1963). One pheromone recently identified is the sex attractant of B. mori. The work on the isolation of this substance was initiated in 1939 and ended with the characterization of this potent volatile material by Butenandt et al. (1959b). This pheromone, given the name bombykol, was identified as hexadec-l0,12-dien-l-ol,a 16-carbon doubly unsaturated alcohol (Butenandt and Hecker, 1961). Jacobson et al. (1960, 1961) have succeeded in identifying the sex attractant of the gypsy Both of these attractants are moth as (d)-lO-acetoxy-7-hexadecen-l-ol. active in unbelievably low quantities and their ultimate action will most likely be measured in numbers of molecules affecting a sense receptor. Nothing has been reported regarding biosynthetic mechanisms or catabolic routes of these substances. 5. Queen substance The queen substance is produced by the mandibular glands of queen bees and acts to prevent gonad maturation in worker bees as well as affect their behaviour. The active pheromone has been shown to be 9-oxodec-trans-2-enoic acid (Barbier and Lederer, 1960; Butler et al., 1961). It is likely that the physiological responses to the queen substance are due to the combined action of the above compound and a yet unidentified compound (Pain et al., 1962). VII. CONCLUSION Although several problems of the function and metabolism of.lipid in insects have been unravelled in recent years, this research area remains ripe for invasion by both the entomologist and biochemist. The task of assuring that insects utilize the same metabolic pathways as micro-organisms and vertebrates has not yet been completed. In this '' pursuit we may uncover new pathways as will most likely be the case for the biosynthesis and catabolism of ecdysone. In the future, gas chromatographic procedures and infra-red spectroscopy will be employed to

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a greater extent and will allow a detailed chemical dissection of insect lipids. Metabolic studies will depart from the isotope injection techniques and will commonly employ in vitro procedures. This will ultimately allow us to elucidate the main sites of lipid synthesis and catabolism on a tissue or cellular basis. Let us hope that more investigators will enter this most fertile area of research and that this review will become rapidly outdated.

ACKNOWLEDGEMENTS Original work from the author's laboratory was supported by grants AM-02818 from NIH and G-22884 and B-17305 from NSF. This article was written while the author held NSF senior post-doctoral fellowship 54023. Sincere thanks are given to the NSF and to Prof. Martin Liischer, Zoologisches Institiit, Universitat, Bern, Switzerland, who provided facilities for conducting some of the research herein reported. Finally the author thanks his students and associates who have worked with him in this research area including Drs H. Chino, K. A. Domroese, R. D. Goodfellow, M. Habibulla and A. W. Wiens as well as Professors H. A . Schneiderman and A. Meyer of Western Reserve University, with whom he has collaborated on the chemistry of the juvenile hormone. REFERENCES

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Weaver, N. and Law, J. H. (1960). Heterogeneity of fatty acids from royal jelly. Nature, Lond. 188, 938-939. Weaver, N., Law, J. H. and Johnston, N. C. (1964). Studies on the lipids of royal jelly. Biochim. biophys. Acta 84, 305-3 15. Weinland, E. (1908). Uber die Bildung von Fett aus eiweissartiger Substanz im Brei der Calliphora Larven. Z. Biol. 51, 197-278. Weis-Fogh, T. (1952). Fat combustion and metabolic rate of flying locusts (Schistocera gregaria Forskal). Phil. Trans. R. SOC.Lond. 237, 1-36. Weiss, S. B. and Kennedy, E. P. (1956). The enzymatic synthesis of triglycerides. J. Am. chem. SOC.78, 3550. Weiss, S. B., Kennedy, E. P. and Kiyasu, J. Y.(1960). The enzymatic synthesis of triglycerides. J. biol. Chem. 235, 4 W . Wigglesworth, V. B. (1928). Digestion in the cockroach. 111. The digestion of proteins and fats. Biochem. J. 22, 150-161. Wigglesworth, V. B. (1942). The storage of protein, fat, glycogen and uric acid in the fat body and other tissues of mosquito larvae. J. exp. Biol. 19, 56-77. Wigglesworth, V. B. (1947). The epicuticle of an insect, Rhodinus prolixus (Hemiptera). Proc. R. SOC.B. 134, 163-181. Wigglesworth, V. B. (1948). The insect cuticle. Biol. Rev. 23, 408-451. Wigglesworth, V. B. (1958). The distribution of esterase in the nervous system and other tissues of the insect Rhodnius prolixus. Q. J. microsc. Sci. 99, 441-450. Wilhelm, R. C., Schneiderman, H. A. and Daniel, L. J. (1961). The effects of anaerobiosis on the giant silkworms Hyalophora cecropia and Samia Cynthia, with special reference to the accumulation of glycerol and lactic acid. J. Insect Physiol. 7, 273-288. Williams, C. B. (1945). Notes on the fat content of two British migrant moths (Lepidoptera). Proc. R. ent. SOC.Lond. 20, 6-13. Williams, C. M. (1956). The juvenile hormone of insects. Nature, Lond. 178, 212-21 3. Williams, C. M. and Law, J. H. (1965). The juvenile hormone. IV. Its extraction, assay and purification. J. Insect Physiol. 11, 569-580. Wlodawer, P. (1956). Studies on the biochemistry of waxmoth. 13. Role of phospholipids in the utilization of wax. Acta Biol. exp. Vars. 17, 221230. Wlodawer, P. and Wisniewoska, A. (1965). Lipids in the haemolymph of waxmoth larvae during starvation. J. Insect Physiol. 11, 11-20. Wojtczak, L., Wlodawer, P. and Zborowski, J. (1963). Adenosine triphosphateinduced contraction of rat liver mitochondria and synthesis of mitochondria1 phospholipids. Biochim. biophys. Acta 70, 290-305. Wolfe, L. S. (1964). Cell membrane constituents concerned with transport mechanisms. Can. J. biochem. Physiol. 42, 971-988. Wootton, J. M. and Wright, L. D. (1960). Biosynthesis of squalene by the annelid Lumbricus terrestris. Nature, Lond. 187, 1027-1028. Wren, J. J. and Mitchell, H. K. (1959). Extraction methods and an investigation of Drosophila lipids. J. biol. Chem. 234, 2823-2828. Wright, L. D. (1961). Biosynthesis of isoprenoid compounds. A. Rev. Biochem. 30, 525-548.

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Wright, L. D. and Cleland, M. (1957). Factors influencing incorporation of mevalonic acid into cholesterol by rat liver homogenates. Proc. SOC.epx. Biol. Med. 96, 219-224. Wyatt, G. R. (1961). The biochemistry of insect haemolymph. A. Rev. Ent. 6, 75-1 02. Wyatt, G . R. and Kalf, G. F. (1957). The chemistry of insect haemolymph. 11. Trehalose and other carbohydrates. J. gen. Physiol. 40, 833-847. Wyatt, G. R. and Meyer, W. L. (1959). The chemistry of insect haemolymph. 111. Glycerol. J. gen. Physiol. 42, 1005-1011. Wyatt, G. R., Kropf, R. B. and Carey, F. G. (1963). The chemistry of insect haemolymph. -1V. Acid-soluble phosphates. J. Insect Physiol. 9, 137152. Yamafuji, I. and Yonezawa, Y. (1935). Enzymes of Bombyx mori. XI. Lipase of the stomach juice. J. agric. Chem. SOC.Japan 11, 77-85. Yamafuji, K. (1937). I. Allgemeine Morphologie des Seidenspinners (Bombyx mori). Tabul. biol. pp, 36-50. Yamamoto, R. T. and Jacobson, M. (1962). Juvenile hormone activity of isomers of farnesol. Nature, Lond. 196, 908-909. Young, R. C. (1963). The biosynthesis of beeswax. Life Sci. 9, 676-679. Young, R. C. (1964a). Digestion of wax by the greater wax moth, Galleria, mellonella. Ann. ent. SOC.Am. 57, 325-327. Young, R. C. (1964b). Lipids of the larva of the greater wax moth, Galleria mellonella. Ann. ent. SOC.Am. 57, 321-324. Zandee, D. I. (1962). Lipid metabolism in Astacus astacus (L.). Nature, Lond. 195, 814-815. Zandee, D. I. (1964). Absence of sterol synthesis in some arthropods. Nature, Lond. 202, 1335-1336. Zebe, E. (1954). uber den Stoffwechsel der Lepidopteren. Z. vergl. Physiol. 36. 290-3 17. Zebe, E. (1959a). Discussion on preceding paper. Niemierko, W. Some aspects of lipid metabolism. Fourth. Znt. Congr. Biochem. 12, 197-199. Zebe, E. (1959b). Die Verteilung von Enzymen des Fettstuffwechsels im Heusschreckenkorper. Verh. d. dtsch. Zool. in Miinsterl West6 309-3 14. Zebe, E. (1960). Condensing enzyme und /3-ketoacylthiolase in verschiedenen Muskeln. Biochem. Z. 332, 328-332. Zebe, E. and McShan, W. H. (1959). Incorporation of 14C-acetateinto long chain fatty acids by the fat body of Prodenia eridania. Biochim. biophys. Acta 31, 5 13-51 8. Zebe, E., Delbruck, A. and Bucher, Th. (1959). uber den glycerin-1 P-cyclus irn Flugmuskel von Locusta migratoria. Biochem. Z . 331, 254-272.

Addenda The following supplementary notes are taken from recent publications. Page 89: The total and free lipid content of the fat body of the ant F. polyctena decreases during the pupal period and the percentage of FFA increases during metamorphosis (Schmidt, 1966). During the embryogenesis of P. americana, the total extractable lipid decreases from 39.5 to 23.2% of the dry weight of the

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ootheca. The greatest decrease was in TGL emphasizing the importance of this class of lipids as an energy source for development (Kinsella and Smyth, 1966). Page 96: The fatty acid composition of mosquito TGL is not affected by temperature (Van Handel, 1966). In Drosophila, diet strongly influences the total lipid content, the quality of lipids and the fatty acid distribution among the lipid classes (Keith, 1966). Thirty different fatty acids from Ca to CS4have been identified in P. americana embryos and the fatty acid patterns suggest that partially hydrolysed TGL molecules are converted to PL during development (Kinsella, 1966a). Page 110: The hemolymph of the waxmoth has the capacity to synthesize TGL from fatty acids. It is postulated that FFA are released from the fat body of this insect rather than DGL (Wlodawer e f af., 1966) but this phenomenon is most likely a form of adaptation of this very specialized insect and is not typical of several other insects studied. Page 131: Calliphora larvae appear to synthesize unsaturated fatty acids via a pathway independent of that used for the synthesis of saturated fatty acids. Palmitate is the major saturated and palmitoleate and oleate the major unsaturated fatty acids (Miura et al., 1965). Studies on A. grandis, however, indicate that there are not two independent pathways for the synthesis of saturated and unsaturated fatty acids. Dietary saturated fatty acids were elongated and/or desaturated by this insect (Lambremont et al., 1965). Page 144: PTE is the major PL constituent of fat body from S. bullata larvae while sphingosine containing lipids appear to be absent (Allen and Newburgh, 1965). PTC and PTE are the major PL fractions in T. molitor larvae (Kamienski et al., 1965). The family Cecidomyiidae of the Diptera differ from other families in having much less palmitoleate and containing more choline glycerophosphatide than ethanolamine glycerophosphatide (Fast, 1965). [P3a]Phosphorylcholinewas incorporated into PTC by the larval fat body of S. bullata at a rate of 4+7%/hrat 30°C (Crone et al., 1966). The qualitative composition of PL did not change during the embryogenesis of P. americana but did increase four fold during this period (Kinsella, 1966b). Page 150: Further work has been completed on the isocitric lyase of Prodenia (Porter and Jaworski, 1965). The conversion of protein and sugar to TGL in the mosquito may occur through a common rate-limiting precursor (Van Handel, 1965). Page 157: Recent reviews on the biosynthesis of sterols include those by Clayton (1965a, b) and Olson (1965). A review on steroids in insects has also appeared (Bergmann, 1965). Page 160: A dietary sterol is necessary for the survival of adult A. grandis (Earle et al., 1965). Page 161: The reproductive activity of well fed desert locusts is hastened and synchronized by terpenes taken in the diet (Carlisle et al., 1965). Page 169: Insects may synthesize terpenes from simpler precursors, as shown by the studies on the synthesis of the defensive secretion by the walking stick, A. buprestoides (Meinwald et al., 1966). Insect embryos cannot synthesize sterols although they can actively metabolize the fatty acids of sterol esters (Kinsella, 1966~). Page 176: Different subcellular membranes of some insect tissues contain similar repeating structural units in which individual sterols occupy sterically characteristic spaces (Lasser and Clayton, 1966). The function of sterol in insects

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is primarily as a component of subcellular membranes. The fat body of a growing insect stores sterols (apparently as esters) that have been displaced from other tissues (Lasser et al., 1966). Pages 179, 180: At least two synthetic substances have been shown to possess potent juvenile hormone activity (Bowers et ul., 1965; Law et al., 1966). Ecdysone has been synthesized by organic chemists (Siddal et al., 1966; Herb et al., 1966). Chemical insect attractants and repellents have been recently reviewed (Jacobson, 1966). One insect has captured a vertebrate hormone as a defensive secretion (Schildnecht et al., 1966). The water beetle stores up to 0.4 mg of cortisone, an amount equal to that extractable from 1,000 ox suprarenal glands.

REFERENCES Allen, R.R. and Newburgh, R. W. (1965). Phospholipid composition of fat-bodies of Surcophaga bullata. J. Insect Physiol. 11, 1601-1603. Bergmann, E. D. (1965). Les steroids des insectes. Bull. SOC.chim., Fr. 2687-2691. Bowers, W. S., Thompson, M. J. and Uebel, E. G. (1965). Juvenile and gonadotropic hormone activity of 10,ll-epoxyfarnesenic acid methyl ester. Lire Sciences, 4, 2323-2331. Carlisle, D. B., Ellis, P. E. and Betts, E. (1965). The influence of aromatic shrubs on sexual maturation in the desert locust, Schistocerca gregaria. J. Insect Physiol. 11, 1541-1558. Clayton, R. B. (1965a). Biosynthesis of sterols, steroids and terpenoids. Part I. Biogenesis of cholesterol and the fundamental steps in terpenoid biosynthesis. Q . Rev. 19, 168-200. Clayton, R. B. (1965b). Biosynthesis of sterols, steroids and terpenoids. Part 11. Phytosterols, terpenes and the physiologically active steroids. Q. Rev. 19, 201-230. Crone, H. D., Newburgh, R. W. and Mezei, C. (1966). The larval fat body of Sarcophuga bullata (Diptera) as a system for studying phospholipid synthesis. J. Insect Physiol. 12, 619-624. Earle, N. E., Walker, A. B. and Burks, M. L. (1965). Storage and excretion of steroids in the adult boll weevil. Comp. Biochem. Physiol. 16, 277-288. Fast, P. (1965). Phylogenetically anomalous lipid composition of Cecidomyiidae (Diptera). Ann. ent. SOC.Am. 58, 933. Jacobson, M. (1966). Chemical insect attractants and repellents. A. Rev. Ent. 11, 403-422. Kamienski, F. X.,Newburgh, R. W. and Brookes, V. J. (1965). The phospholipid pattern of Tenebrio molitor larvae. J. Insect Physiol. 11, 1533-1540. Keith, A. D. (1966). Analysis of lipids in Drosophila melanogaster. Comp. Biochem. Physiol. 17, 1127-1136. Herb, U., Hocks, P., Wiechert, R., Furlenmeier, A., Furst, A., Langemann, A. and Waldvogel, G. (1966). Die Synthese des Ecdysons. Tetrahedron Lett. 13, 1387-1 391. Kinsella, J. E. (1966a). Metabolic patterns of the fatty acids of Periplaneta americana (L.) during its embryonic development. Can. J. Biochem. 44,247258. Kinsella, J. E. (1966b). Phospholipid patterns of Periplaneta americana during embryogenesis. Comp. Biochem. Physiol. 17, 635-640.

ADDENDA

21 1

Kinsella, J. E. (1966c). Sterol metabolism during embryogenesis of Periplaneta americana (L.). J. Insect Physiol. 12, 435-438. Kinsella, J. E. and Smyth jr., T. (1966). Lipid metabolism of P. americana during embryogenesis. Comp. Biochem. Physiol. 17, 237-244. Lambremont, E. N., Stein, C. I. and Bennett, A. F. (1965). Synthesis and metabolic conversion of fatty acids by the larval boll weevil. Comp. Biochem. Physiol. 16, 289-302. Lasser, N. L. and Clayton, R. B. (1966). The intracellular distribution of sterols in Eurycofis floridana and its possible relation to subcellular membrane structure. J. Lipid Res. 7, 413-421. Lasser, N. L., Edwards, A. M. and Clayton, R. B. (1966) Distribution and dynamic state of sterols and steroids in the tissues of an insect, the roach Eurycotis floridana. J. Lipid Res. 7, 403-412. Law, J. H., Yuan, C. and Williams, C. M. (1966). Synthesis of a material with high juvenile hormone activity. Proc. natn. Acad. Sci. U.S.A. 55, 576-578. Meinwald, J., Happ, G. M., Labows, J. and Eisner, T. (1966). Cyclopentanoid terpene biosynthesis in a phasmid insect and in catmint. Science, N . Y. 151, 79-80. Miura, K., Vonk, H. J. and Zandee, D. I. (1965). Biosynthesis of unsaturated and saturated fatty acids in aspectically reared larvae of Calliphora erythrocephala (Meig.). Archs int. Physiol. Biochim. 73, 65-72. Olson, J. A. (1965). The biosynthesis of cholesterol. Ergebn. Physiol. 56, 173-215. Porter, C. A. and Jaworski, E. G. (1965). Biosynthesis of chitin during various stages in the metamorphosis of Prodenia eridania. J. Insect Physiol. 11, 11511160. Schildnecht, H., Siewerdt, R. and Maschwitz, U. (1966). A vertebrate hormone as defensive substance of the water beetle Dytiscus marginalis. Angew. Chem. 5, 421-422. Schmidt, G. H. (1966). Probleme der Fettmobilisierung bei Insekten. I. Veranderungen der Freien und Gebundenen Lipide, des Lipoid-Phosphors sowie der Freien und Veresterten Fettsauren wahrend der Pupaphase von Formica polyctena. J. Insect Physiol. 12, 237-249. Siddall, J. B., Cross, A. D. and Fried, J. H. (1966). Steroids ccxcii. Synthetic studies on insect hormones 11. The synthesis of ecdysone. J. Am. chem. SOC. 88, 862-863. Van Handel, E. (1965). The obese mosquito. J . Physiol. 181, 478-486. Van Handel, E. (1966). Temperature independence of the composition of triglyceride fatty acids synthesized de novo by the mosquito. J. Lipid Res. 7, 1 1 2-1 1 5. Wlodawer, P., Lagwinska, E. and Baranska, J. (1966). Esterification of fatty acids in the wax moth haemolymph and its possible role in lipid transport. J. Insect Physiol. 12, 547-560.

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Chitin Orientation in Cuticle and its Control A. C . NEVILLE

ARC Unit of Muscular Mechanisms and Insect Physiology, Department of Zoology, University of Oxford, England 213 I. Introduction . 215 A. Locust Solid and Rubber-like Cuticles . 11. Orientation and the Mechanical Properties of Cuticular Macro217 molecules . 220 111. Types of Chitin Architecture . 220 A. Parallel orientation . 220 B. Crossed fibrillar orientation . 223 C. Lamellar structure . 229 D. Functional aspects . -233 IV. Orientation Control . 233 A. Circadian organization. . 246 B. Metabolic oscillators and “switches” . 254 C. “Dermal” light sense . 257 D. Implantation experiments . , . 260 E. Nervous control . 260 F. Discussion . 262 V. Orientation Mechanisms . 263 A. Primary orientation . 265 B. Secondary orientation . 270 C. Protein orientation . 272 D. Hypothesis . 279 VI. Conclusion . 280 References . 286 Addenda .

I. I N T R 0D U CT I 0 N One aspect of the study of differentiation concerns the way in which macromolecular polymers become oriented in skeletons and cell walls. This aspect is an interdisciplinary problem, involving many kinds of biological macromolecules and a variety of plants and animals. Recent experimental work on the orientation of chitin in insect cuticle has begun to reveal a complex of physiological factors which affect the final architectural product. This article deals largely with this field and also incorporates some unpublished material. Several structural polysaccharides (chitin, cellulose, tunicin) show great similarity at the 213

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macromolecular level and, where it has appeared fruitful, comparison has been made with chitin orientations in other arthropods, and also with the various other fibrillar systems produced by plants, bacteria, vertebrates and tunicates. In uitro reconstitution experiments with collagen (Randall et ul., 1955), with bacterial flagellin (Asakura et ul., 1966) and with myosin (Huxley, 1963), indicate that the problem of macromolecular orientation in organic systems may ultimately be solved at the physicochemical level. This would suggest that the biologist’s task is to concentrate upon the relevant control systems which are involved. For example, the fact that microfibrils occur in synthetic polymers shows that they can form without the agency of living protoplasm (Ribi, 1951). Further, the apparent control of microfibril diameter is clearly not a phenomenon specifically associated with living systems (Table I). TABLE I Some organic and synthetic microfibrils and their diameters Source

Microfibril diameter

Reference

Plants and bacteria

25M00 A

Preston et al. (1948) Frey-Wyssling et al. (1948) Miihlethaler (1950) refs. in Ribi (1951) Ohad et al. (1962) Meyer et al. (195 1) Ribi (1951) Locke (1961) Richards and Korda (1948) McLachlan et al. (1965)

Plant cellulose Bacterial cellulose Tunicate cellulose Chitin, Crustacea Chitin, insect endocuticle Chitin, insect tracheae Chitin, diatom, extracellular and without protein Viscose rayon, reprecipitated cellulose Polyamides (e.g. c-amino caproic acid) Polyethelene plastic

100 A 16 8,x 30 A 200 A 100 A 25 8, 75-300 A 20C300 A 100 A

Ribi (1951)

100 A

Ribi (1951)

loo A

Ribi (1951)

However, the type of orientation known as “crossed fibrillar” (see below) appears to date to be a phenomenon without parallel in the process of inorganic crystallization. Microfibril synthesis and microfibril orientation clearly involve different mechanisms as is shown, firstly, by the synthetic microfibrils in Table I; secondly, by chitin

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215

orientation changes which can be induced during the deposition of insect cuticle (Neville, 1963d, 1965a, b, c, d); and thirdly, by cellulose orientation changes in plant cell walls, either natural or induced (Preston, 1961). In the present review it will be established that the insect epidermal cell has control of the extracellular environment in which orientation occurs, although the quantitative contributions made by cytoplasmic control and natural aggregatory properties of the organic polymers is not yet known. A general scheme of events leading to the orientation of chitin is given below. It is an expansion of the schemes of Colvin (1964), for cellulose in plant cell walls, and of Ben-Hayyim and Ohad (1965), for cellulose outside bacterial cell walls : (a) polymerization of the activated monomeric precursor to form a high M molecule (/3 1-4 covalent linkage formation). (b) transport of the molecule from the site of synthesis to that of crystallization, if these are different. (c) crystallization or fibril formation (H-bonding). (d) primary orientation of fibrils during deposition together with its control mechanism. (e) secondary reorientation of fibrils together with its control mechanisms. The distinction between primary orientation and secondary reorientation events is described below. It is sufficient to note here that their control systems can be separated experimentally in the deposition of locust solid cuticle. Since much of the experimental work to be reviewed concerns locust solid and rubber-like cuticles, the distinctions between them are given here. A. LOCUST SOLID AND RUBBER-LIKE CUTICLES

These two kinds of cuticle were first distinguished on the basis of their physical and chemical properties by Weis-Fogh (1 960). Solid cuticle is a two-phase solid with a mean value for Young’s modulus of 9.4~ 1O1O dyn/cm2 (Jensen and Weis-Fogh, 1962), a value which, for similar stresses, lies between those of its two main components, the high modulus polysaccharide chitin and the lower modulus cuticular proteins. (Cuticle does not obey Hook’s law, so that Young’s modulus varies with stress.) Rubber-like cuticle, on the other hand, has a Young’s modulus of approximately 2 x lo7 dyn/cm2, which is typical of elastomers (Jensen and Weis-Fogh, 1962). The remarkable elastic 9 + A.I.P. 4

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properties of locust rubber-like cuticle are due to the protein resilin, whose properties have been described in the review of Andersen and Weis-Fogh (1964). Some samples of locust rubber-like cuticle also contain chitin, and are thus convenient places to study its deposition (Neville, 1963b), since the only other component present is resilin itself. This review is concerned with the orientation of naturally occurring chitin-protein complexes and not with the artificially reoriented chitin which results from deproteinization procedures.

10-28H

FIG. I . Eight residues of a-chitin (carbon atoms drawn as closed circles; oxygen as open circles; nitrogen closed circles with rings; hydrogen bonds are drawn as dotted lines). The 31 A repeat along the chains, probably caused by the absence of some acetyl groups (Rudall, 1963), is indicated by the dotted circles. (After Carlstrom, 1957.)

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11. O R I E N T A T I OAN N D T H E M E C H A N I C APL R O P E R T I EOSF

C U T I C U L AM RA C R O M O L E C U L E S Chitin is a structural polysaccharide which is physically and chemically anisotropic. This is the fundamental basis for the evolution of the many kinds of molecular architecture displayed by native chitin in biological material. The physical properties of chitin are primarily determined by its macromolecular configuration (@-1,4linked polymer chains) : similarly the configurations of cuticular proteins determine their basic mechanical properties, e.g. @-extended “arthropodins” (mechanically anisotropic), and randomly kinked, randomly oriented polypeptide chains of resilin (mechanically isotropic). The particular value of the /3-I, 4 link is well discussed by Preston (1964) with regard to the similar linkage in plant celluloses. The twofold screw axis with alternation of side chains (Fig. 1) means that the chains are straight, as opposed to the helices of a-I, 4 linked starch and /I-1, 3 linked xylan.* Helical chains, even when they form microfibrils (e.g. in xylan, Frei and Preston, 1963), do not possess the particular advantage of /3-1,4 linked polymers, namely, the possibility that all of the covalent backbone linkages can be oriented parallel to the microfibril axis. The prime use of well oriented backbone linkages is in improved tensile strength of materials. With a lesser degree of orientation there is a smaller number of molecular chains simultaneously under tension and hence a greater stress concentration upon them. This effect lowers the tensile strength since the most heavily loaded chains rupture first. Since chitin is the chief tensile component of insect cuticle we would expect to find it maximally oriented in those parts of insect skeletons which are subjected to the greatest stresses (e.2. wing veins, jumping legs, ovipositors, muscle tendons); X-ray diffraction (Rudall, 1963) and polarization analysis (Neville, 1965b) confirm this. Results from X-ray diffraction and infra-red spectroscopy (Carlstrom, 1957) show that, in samples of chitin in which the molecular chains run parallel, the interchain bonding is as shown in Fig. 2. The b axis is the strongest direction (covalent backbone links and intrachain hydrogen bonds). The next strongest is along the a axis (hydrogen bonding). The a and b axes run parallel to the cuticle surface in blowfly larval and pupal cuticle (Fraenkel and Rudall, 1947). Hence the weakest direction is the c axis (van der Waal’s forces) which runs perpendicular to the cuticle surface; this provides one molecular basis for the laminate * A helicoidal model for cellulose has been proposed (Manley, R. St. John, The fine structure of cellulose microfibrils, Nature, Lond. 204, 1155-1 157,1964: Manley, R. St. John and Inoue, S., The fine structure of regenerated cellulose, J . Po/vtner Sri. B3,691-695, 1965).

218 A . C. NEVILLE cleavage of cuticle parallel to its surface. The disadvantage of an arrangement which confers strength by orienting all of the covalent links in one particular direction (e.g. along the axis of a wing vein, leg, ovipositor or muscle tendon), is that the structure is weak when twisted, which causes a shearing of weaker bonds. Parallel orientation is effective, however, in muscle tendons which transmit force axially without a twisting component, but would be disadvantageous in the

FIG.2. A block diagram of a piece of locust leg cuticle in which most of the chitin chains are oriented along the leg as indicated. The arrows indicate strength in various directions ( b axis: covalent bonds plus hydrogen bonds; a axis: hydrogen bonds; c axis: van der Waal's bonds).

other structures. There is thus a mechanical advantage in the growth of cuticular layers with periodic changes in orientation. Experimental interference with cuticular morphogenesis weakens the final structure (Neville, 1965b, and below). It is worth noting here that solid cuticle, bonded exclusively by covalent linkages, would be too brittle. The presence of weak secondary bonds serves to counteract this. On these grounds, Jensen and Weis-Fogh (1962) have criticized the cuticle model of Fraenkel and Rudall (1947) which proposed covalently bonded alternating monomolecular layers of chitin and protein (Fig. 3). The quantitative aspects of improving strength by orientation are well known in the textile industry. Orientation has an effect on both the stress-strain curve and on the rupture strength. Rupture of a fibre at breaking point may result from fracture of the covalent backbone linkages or from slipping of the chains or crystallites over one another. A fibre which is built up exclusively from long chains and which fails by fracture of the chains, should not be weakened by swelling agents (e.g. water) which reduce the cohesion of chains and micellae. A fibre of shorter chains which fails by slipping will be weakened by water. According to Thor and Henderson (1940) the tensile strength of dry purified chitin is 9.5 kg/mm2 and that of wet purified chitin is 1.8

C H I T I N ORIENTATION I N CUTICLE A N D ITS CONTROL

219

FIG.3. The compatibility of chitin and B-arthropodin chains when oriented in parallel. Every fourth glucosamine residue coincides with every sixth amino acid residue, as indicated by dotted circles (after Fraenkel and Rudall, 1947).

kg/mm2. However, the purification process may partly explain this result which is at variance with the properties of other natural fibres (Table 11). The high value for the balken fibre of a Goliath beetle (Goliuthus) calculated from Herzog’s (1 926) stress-strain curve could, as suggested by Jensen and Weis-Fogh (i962), be due to the use of an unpurified fibre, although this is not known for sure. There is a need TABLEI1 Tensile strength of chitin and some other natural materials Material

Tensile strength (kg/mm2) Dry Wet ~

Ramie Cotton Flax Purified chitin Dry “chitin”

~

.

91-95 20-80 a4 9.5 58

_

_

_

_

108 24-83 88 1.8

_

Reference _

Meyer (1 950) Meyer (1950) Meyer (1950) Thor andnHenderson (1940) Herzog (1926)

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A . C . NEVILLE

for further work on cuticles as two phase systems, along the lines of Currey’s (1 964) analysis for bone. In the past there was controversy as to whether the intrinsic birefringence of chitin is positive or negative. In contrast to cellulose, which has a positive intrinsic birefringence, chitin possesses acetamido side chains which make its intrinsic birefringence negative, since the side chains are oriented almost perpendicularly to the chain axis (Fig. l), (cf. negative birefringence of DNA caused by base pairs oriented perpendicularly to the chain axis). X-ray diffraction analysis shows that naturally occurring chitin displays preferred orientation of crystallites (Clark and Smith, 1936; Carlstrom, 1957; Rudall, 1963). Because it usually occurs in oriented structures, chitin displays a strong form birefringence when immersed in media of suitably different refractive index from that of chitin itself; its form birefringence is usually many times greater than its intrinsic birefringence. The orientation of chitin fibrils can, therefore, be deduced from form birefringence studies carried out in water. 111. T Y P E SO F C H I T I NA R C H I T E C T U R E An exact knowledge of the chitin architecture of localized regions of insect skeletons is necessary for the interpretation of experimental morphogenetical studies, since cellular control of orientation differs from species to species and also from region to region within an insect. The purpose of this section is therefore to provide a few chosen examples of chitin orientation which occur in cuticles. A . PARAL.LEL ORIENTATION

The parallel orientation of chitin chains into fibres to give strength along the axis is widespread. Recognizable fibre tracts cr “balken” are found in such structures as muscle apodemes and tendons, in the endocuticle of wing veins and legs, in ovipositors and in swimming hairs. This type of orientation is a component of the next type. B . CROSSED FIBRILLAR ORIENTATION

In crossed fibrillar structures the molecular chains of any one layer are oriented in parallel, with the direction of orientation changing from layer to layer to give a multi-ply lamellate. Crossed fibrillar structures in animals have been well reviewed by Picken (1960). In insects, crossed fibrillar chitin has been known in beetles for many years (older refs. in

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Richards, 1951). In the present investigations they have been found to occur throughout the skeleton in examples from Elateridae, Scarabaeidae, Buprestidae, Carabidae, and Lucanidae and are probably widespread. The architecture of the hind femur of Heliocopris colossus (Scarabaeidae) is shown in Fig. 4. Alternating equiangular left and right-

FIG.4. Diagram of the alternating equiangular left- and right-handed helices of chitin fibrils (balken) in the hind femur of a beetle Hdiocopris co/ossus (Scarabeidae). Directions of maximum retardation indicated by biaxial indicatrices.

handed helices of chitin fibrils wind around the leg, crossing orthogonally as shown. Transverse sections of the leg show very little difference in orientation from layer to layer as the cut runs at 45" to both fibril directions. Sections cut along one of the fibril directions show alternating birefringent and isotropic layers when viewed between crossed polaroids. The system is reminiscent of the helically wound collagen fibres in Haversian systems of bone (Gebhardt, 1901). A similar system of crossed fibrillar chitin has been found in the legs and wings of toebiter water bugs (Hemiptera : Belostomatidae, Neville, 1965d). It occurs in all of the species examined (Hydrocyrius colombiue, Lethocerus cordofanus and L. uhleri, and Lirnnogeton,fieberi). The formation of the crossed fibrillar chitin of Belostomatids has been found to be controlled by a circadian clock, a sharp change in angle (varying from 30" to 90" from region to region) of orientation occurring every 24 h (Neville, 1965d). This sharp transition in orientation contrasts with the gradual orientation change proposed to explain the birefringent properties of lobster cuticle (Schmidt, 1924). Schmidt was unable to cut sections in which the lamellae all appeared the same in the polarizing microscope, which showed that they were not orthogonally arranged as Biedermann (1903) had suggested. This problem still remains unsolved, but a system like the Belostomatid cuticle with the direction of orientation changing by an angle differing from 90" could provide a solution. It is not yet

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known whether the crossed fibrillar system of beetle cuticle is circadially controlled, but the appearance of sections in the polarizing microscope indicates that this is likely. The system seems to be associated with extra strengthening of the cuticle in large beetles, which would seem particularly favourable material for further work. The crossed fibrillar chitin structure described in honey bee (&is mellqera) antennal cuticle (Richards, 1952) is clearly not coupled to a circadian mechanism (Fig. 5). In this case the endocuticle is all grown

FIG.5. Diagram of a surface view of the crossed fibrillar chitin fibrils in the antennal cuticle of the honey bee Apis mellifica. (After Richards, 1952.)

in a single day and the 15 to 20 fibril direction changes all occur in the course of the one day (see Section 1V B). Other examples of crossed fibrillar chitin in insects are the peritrophic membrane (Mercer and Day, 1952) which appears to be laid down in a template of microvilli; the crossed fibre pattern of butterfly scales (Anderson and Richards,

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1942); and the system of taenidia in insect tracheae, whose orientation has been interpreted in terms of forces resulting from expansion and buckling of the cuticulin (Locke, 1958, 1964). C . LAMELLAR S T R U C T U R E

All arthropod cuticles are laminate in texture. This is the expected product of a single layered epithelium, whose cells have a marked tendency for adhering laterally, so as to present a surface of contact with the internal and external media (Picken, 1960). Hence, when epithelia secrete external skeletons these are usually laminated in planes parallel to the skeletogenic cells. Lamellae are found in the solid cuticle which forms the bulk of the arthropod skeleton, in the rubberlike cuticle of the elastic ligaments of the flight system (Weis-Fogh, 1960; Neville, 1963a), and in the soft arthrodial membranes (e.g. Malek, 1958). One common appearance of cuticular lamellae is drawn in Fig. 6. They may be referred to as parabolic lamellae. A table of occurrence for such lamellae is given in Table 111. It can be seen that this basic pattern occurs over a wide range of thicknesses (0.1 to 50 microns [p]); in egg, larva, pupa and adult; in solid cuticle and arthrodial membrane; and in all the main classes of arthropods. The pattern also occurs in other biological systems (e.g. bone, dinoflagellate chromosomes), and is briefly reviewed by Bouligand (1965b). To these may be added its occurrence in the tunicin of tunicates (A. C . Neville, unpublished) and collagen in the vertebrate cornea (Jakus, 1964). Rudall (1963) produced X-ray diffraction evidence for parabolic lamellae in blowfly puparium cuticle. The oval diffraction ring recorded indicates that the chitin chains are spaced closest together parallel to the cuticle surface, and furthest apart in fibrils perpendicular to the surface, the spacing increasing progressively from the former to the latter orientation. Such evidence agrees with microscopical observations, and also with the fact that the density of the parallel region is higher than that of the rest of a lamella. The widespread occurrence of the parabolic pattern suggests that there may be some fundamental process which determines its manner of organization. It is interesting that the direction of the parabolae is polarized in all of the recorded examples. Locke (1964) notes that parabolic lamellae are most easily resolved in the electron microscope in the plastic cuticles of caterpillars, and suggests from model experiments that it ‘is a device which, when combined with few cross-links, can be easily distorted in three dimensions. g*

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The parabolic lamellae of the tunicate Cynthiu papillosa, first described and illustrated as laminae with feathery margins by Schultz (1863), are so wide (50 p) that they can be manipulated and observed under the polarizing light microscope (A. C . Neville, unpublished). The

FIG.6. Diagram of the apparent microfibril directions in two parabolic larnellae seen in section (after Locke, 1961).

parabolic fibrils are compressed or extended perpendicular to the lamellar surface, when the sample is compressed or extended. Observations between crossed polaroids and in ordinary light and phase contrast show the parallel region to be the densest part of a lamella. The course of tunicin fibrils from one lamella to the next was traced in optical

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sections by focusing up and down on a slice of test. The result agreed with the suggestion of Drach (1953) for the course of the chitin fibrils in the lamellae of lobster (Homarus) cuticle. The parabolae lie in a plane which is not perpendicular to the lamellar surface, but is inclined to it at an angle (Fig. 7). This results in the appearance of parabolae in any section taken perpendicularly or horizontally through the cuticle or test, and explains the sequence of directions of parabolae observed when a rectangular slab of tunicate test was rotated under the polarizing microscope (left, left, right, right). Slifer and Sekhon (1963) found parabolic microfibrils in sections of Melanoplus egg shells taken obliquely to the cuticle surface.

FIG.7. The course of parabolic fibrils of tunicin in two adjacent lamellae from the test of a tunicate, Cynthiu yapillmu. The top faces of the cubes run parallel to the test surface. Similar deductions have been made for the course of chitin fibrils in the cuticle of a lobster, Homarits sp. (Drach, 1953.) Scale: cubes are drawn with 5 0 p sides.

There is at present a controversy as to whether parabolic fibrils as such exist, or whether they are the result of MoirC patterns formed from fibrils seen in oblique sections. The latter view has been proposed for the cuticle of some Crustaceans by Bouligand (1965a), who has examined the Copepod Acanrhocyclops viridis (Jurine), and the crabs Carcinus moenas L., Cancer yagurus L. and Macropipus puber L. Bouligand’s interpretation depends upon parallel orientation plane to the cuticle surface in each submicroscopically thin layer, combined with a small progressive change in angle of orientation from layer to layer (Fig. 8).

TABLE 111 The occurrence of parabolic lamellae in arthropods and tunicates Material

Stage

Calpodes (Lepidoptera) Calpodes Galleria mellonella (Lepidoptera) Tenebsio niolitor (Coleoptera)

Pupa Larva Larva Larva

Tenebsio molitor Apis niellifera (Hymenoptera) Melanoplus diflerentialis (Orthoptera)

Pupa Adult

Melanoplus difesentialis

Exuvia

Approx. spacing of lamellae

Reference

Notes

Locke (1961) Locke (1 960) Locke (1961)

As in Fig. 6 As in Fig. 6 As in Fig. 6

2.0 p

Locke (1961)

1.0 p 1.0 p

Locke (1961) Locke (1961)

Not fibrous, but pore canals arranged in similar pattern in oblique sections As in Fig. 6 As in Fig. 6

0.3 p 0.1 to 1.0 p 0.5 to 1.0 p

z

m

5

r r

n

Egg she!l

Slifer and Sekhon (1963)

0.75 p

Taylor and Richards (1965)

Embryonic endocuticle secreted by serosa, forming part of egg shell. Pattern as for Fig. 6, but plane to cuticle surface As in Fig. 6, ecdysial membrane

0.2 to 0.3 p

Podura aquatica (Collem bola) Isoptera

Adult

Orconectes viridis (Crustacea)

Adult

2.5 to 7.5 p

Orconectes viridis Orconectes viridis

Adult Adult

5.0 to 16 p 0.5 p

Astacus astacus (Crustacea) Astacus astacus

Adult

20 tL

Homarus (Crustacea) Rhinocricus nodulipes (Diplopoda) Limulus (Arachnida)

Adult Adult

Cynthia papillosa (Tunicata)

Adult

Adult

Adult

Noble-Nesbitt (1963a)

Arthrodial membrane at base of setae Noirot (pers. comm.) As in Fig. 6, arthrodial membrane of physogastrous female Travis and Friberg (1963) Exocuticle, as in Fig. 6, but with fibres branching not only up and down but also sideways Travis and Friberg (1963) Endocuticle, as above Travis and Friberg (1 963) Membranous layer (innermost endocuticle), as above Solid leg endocuticle, as in Fig. 6 Neville (1965 b) Dennel (1960)

10 P

Drach (1 953) Silvestri (1 903) Neville (unpub.) Schulze (1 863) Neville (unpub.)

Fibres less well ordered than in Fig. 6 As in Fig. 6 As in Fig. 6 Coxo-trochanteral arthrodial membrane. Fibres less well ordered than in Fig. 6 Fibrils of tunicin, crystallographically similar to Cellulose I11

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A . C. N E V I L L E

FIG.8. A diagrammatic representation of the possible derivation of parabolic patterns in cuticle. A truncated pyramid of cuticle is viewed from the surface. The direction of orientation of chitin molecules is parallel in each sub-microscopically thin layer, and the direction changes from layer to layer. Oblique sections (faces of the pyramid) then produce Moire patterns with apparent parabolic fibrils, four of which are indicated. (After Bouligand, 1965a.)

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This is essentially the explanation offered by Schmidt (1924) to explain the structure of lobster cuticle. Such a structure would provide a unified basis to explain cuticular stratifications on several size levels, based simply upon the number of orientation changes per quantity of cuticle secreted. Thus in the examples quoted by Bouligand there would be many orientation changes per lamella (probably some hundreds per micron thickness of cuticle) ; in honey bee antenna1 cuticle, each direction change appears as a lamella of which 15 to 20 are constructed in a single day (giving approximately one direction change per micron thickness of cuticle) (Richards, 1952); finally, in Belostomatid water bugs a 5 p thick daily growth layer results from a single orientation change per day (Neville, 1965d). It is possible that both parabolic fibrils and Moire effects occur in different systems. Bouligand’s model (Fig. 8) does not fit the tunicate case for two reasons. (1) The alignment of parabolae on consecutive faces of the pyramid model changes by half a register. This does not occur in tunicate lamellae. (2) The direction of parabolae in the model does not change from face to face, whereas in the tunicate example (as can be worked out from Fig. 7) the sequence is left, left, right, right. The fine structure of the lamellae of locust and cockroach solid cuticle, in which experimental investigations of chitin lamellogenesis are reviewed below (Section IV), has not yet been established. The time is ripe for an electron microscope study of orthopteran cuticle and indeed for a study of chitinogenic cells in general. D . FUNCTIONAL ASPECTS

The prime function of chitin orientation is for improved strength in certain directions. Examples are legion, and only one or two representatives will be mentioned here. Evidence that chitin lamellae in locust endocuticle have some effect upon structural stability has been obtained by comparing the swelling in 3% acetic acid of deproteinized samples of lamellate and non-lamellate cuticle, prepared as described in Section IV A (Neville, 1965b). The framework of the non-lamellate sample was swollen and dispersed more quickly than that of the lamellate sample (Fig. 9). Further evidence for the stabilizing influence of lamellae in locust exocuticle is given in Section V B. In belostomatid bugs there are pits running through the cuticle to the base of tactile spines (Fig. 10). The pits are strengthened around the

FIG.9. Morphogenetically altered locust cuticle. Portions of experimentally produced non-lamellate tibia (top) and lamellate tibia (bottom), initially cut to the same size. The samples were deproteinized and the chitin deacetylated by hot alkali, and swollen in 3% acetic acid. The lamellate sample is more resistant to swelling and dispersion than the non-lamellate sample (from Neville, 1965b).

FIG. 10. Transverse section of hind leg of Hvdroryrius rdombiue (Hemiptera: Belostomatidae), photographed between crossed polaroids. The daily growth layers appear as alternately anjsoptropic and isotropic layers.

FIG. 11. Surface view of two layers of endocuticle cleaved from a leg of Hydrocyrius, and showing crossed fibrillar structure. The rims of the ducts are strengthened by more closely packed and better oriented material.

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FIG.12. Transverse section of hind tibia of Hydrorvrius colombiae (Hemiptera: Belostomatidae), photographed between crossed polaroids to show the strengthening bars of cuticle running across the tibia.

edges by well-oriented closely packed chitin fibrils (Fig. 11) which counteract the stress concentrations occurring at the edges of the holes (Timoshenko, 1951). In regions of closely opposed cuticle (e.g. elytra, swimming legs) in some beetles and belostomatid bugs, cuticular connections dowel the two sheets of cuticle together across the intervening space. The chitin orientation is parallel turning through a right angle to cross the intervening space (Fig. 12). In the beetle Sternoceru castanea Oliv. (Buprestidae) the dowelling pegs connecting dorsal and ventral surfaces of the elytra are strengthened even further by being orthogonally plied in layers, the chitin spiralling alternately in left- and right-handed helices in consecutive concentric layers of the peg. A good example of a strategically oriented structure is the scalariform strut developed on the inside walls of the pleura of the dragonfly thorax. The struts function in providing thoracic elasticity for the flight system and are only developed in strong fliers. The orientations of the chitin fibrils are shown in Fig. 13. As a final example, the cleavage direction of dipterous puparia at adult emergence is determined by the orientation of chitin chains (Section V B).

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FIG. 13. A diagram of the scalariform apodeme which strengthens the thoracic wall of the adult dragonfly, Aeslina jrincea (Odonata: Aeshnidae). The directions of orientation of chitin fibrils are indicated by biaxial indicatrices.

LV. O R I E N T A T I O CN ONTROL This section will be concerned with the interactions of internal and external factors, the interplay of which may be superimposed upon the basic chitin orientation mechanism dealt with in Section V below. It is worth remembering that it would not be surprising to find a different manner of control in each insect order, a situation paralleling their flight control systems. A.

CIRCADIAN ORGANIZATION

The discovery of daily growth layers in the endocuticle of several insects (Neville, 1963c, d) has led to experiments establishing the involvement of a circadian clock in chitin orientation control (Neville, 1965d). The results provide good evidence that chitin orientation is under cellular control. A list of insects in which the presence of daily growth layers has now been established is given in Table IV. Daily growth layers are found in both nymphs and adults, occurring in tibiae, femora, thorax, wing veins, ovipositors, ocelli and compound eyes. They appear to be a general characteristic of cuticle growth in arthropods, occurring in the following systems : (a) Daily rhythm of chitin lamellogenesis in locust and cockroach solid endocuticle, known to be circadian (Neville, 1965d).

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A . C. NEVILLE

(b) Daily change in chitin crossed fibrillar directions in belostomatid bugs, known to be circadian (Neville, 1965d). The same effect may also exist in beetles. (c) Daily fluorescent growth layers in resilin in locust rubber-like cuticle (Neville, 1963a), possibly reflecting daily variation in the extent of cross-linking of the molecular network (Neville, 1963b) by dityrosine and trityrosine (Andersen, 1964). The daily growth layers of resilin have been confirmed by marking it (at known intervals) with injected tritiated tyrosine, when the incorporated tyrosine appears as discrete bands in subsequent autoradiographs (Kristensen, 1966). The bands appear in predictable sites with respect to the daily growth layers. (d) Daily rhythm of calcification of crustacean gastroliths (Scudamore, 1947). TABLE IV List of insects with daily growth layers caused by chitin orientation changes in the endocuticle (compiled from Neville, 1963a-d, 1965a-c and unpublished) Order

Orthoptera

Dictyoptera Phasmida Dermaptera Odonata Hemiptera-Heteroptera Hemiptera-Homoptera

Species

Circadian clock involved

Decticus uerruciuorus Decticus verrucivorus (nymphs) Schistocerca gregaria Schistocerca paranamerise Locusta migratoria Nomadacris septernfasciata Omocestus viridulus Gomphocerus rnaculatus Dolichopoda linderi Stenobothrus lineatus Tettigonia viridissinia Periplaneta arnericana Periplaneta americana (nymphs) Siphyloidea sp. Forficula auricularia Aeshna juncea Aeshna grandis Oncopeltus fasciatus Hydrocyrius colonibiae Aphrophora alni

? ?

Yes Yes Yes ? ? ?

Yes ? ?

Yes Yes 9

? ? ? ?

Yes ?

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235

Locust cuticle is convenient for experiments on chitin orientation for several reasons. (1) It is transparent except for locally melanized regions of the exocuticle, making possible the analysis of the form birefringence of its various fibril systems in the polarizing light microscope. (2) The chitin fibril systems in the locust hind tibia lie almost orthogonal to each other, simplifying optical interaction between them. (3) The orientation of the structures, and thus their form birefringences, can be changed by varying the rearing conditions at the time of deposition. (4) Different epidermal cells, always constant in location, react in varying ways to specific combinations of environmental factors. 1. The chitin orientations in a locust hind tibia reared in control conditions In the desert locust, Schistocerca gregaria, the deposition of exocuticle and endocuticle is separated by ecdysis. The lamellae of exocuticle are spaced too closely together to be resolvable in the light microscope, but they can be shown to cause the strong positive birefringence parallel to the cuticle which is observable in sections, by treatment in 20% NaOH at 160" for I $ h followed by chitosaniodide staining. Cuticle which loses its protein shrinks, so that if the

FIG. 14. Transverse section of adult locust hind tibia photographed between crossed polaroids, 45" off extinction for maximum brightness of birefringence. This section was taken from an adult reared in control conditions and killed 5 days after emergence. Low magnification shows 5 pairs of daily growth layers in the endocuticle. Scale represents 50 p (from Neville, 1965b),

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A. C. NEVILLL

FIG. 15. High magnification reveals that the pairs of daily growth zones consist of lamellate layers alternating with non-lamellate layers. Each lamellate layer consists of a group of birefringent lamellae (running from top to bottom in the photograph). Birefringent pore canal components run from right to left. The experimental history of the zones photographed is, reading from left to right, 12 h day, 12 h night, 12 h day, 12 h night, 60 h prolonged day inhibiting lamellation. 12 h night, 12 h day, 12 h night; scale represents 10 p (from Neville, 1965b).

sections are then pressed out to beyond their original size, the now purple-stained lamellae become sufficiently separated to be individually resolvable in the light microscope (Neville, 1965b). They have since been observed in the electron microscope (A. C.Neville, unpublished). Locust endocuticle grows by daily increments, chronicling its progress by daily growth layers (Neville, 1963c, d). The zonation is due to differences in diurnal and nocturnal deposits, one pair of layers being deposited every 24 h. Since the diurnal zonation in solid cuticle is due to changes in chitin orientation (see below), it is best observed in the polarizing light microscope. Low power photomicrographs of transverse sections of the solid cuticle of locust legs, viewed between crossed polaroids, show alternate concentric layers with strong and weak birefringence (Fig. 14). Although at low power these appear to be arranged with an orthogonal change in orientation, as in plywood and in the haversian systems of bone, higher magnification (Fig. 15) reveals them to consist of lamellate layers alternating with non-lamellate layers. The lamellae can be shown, both histochemically and chromato-

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graphically, to contain chitin (Neville, 1965b). Although experimental removal of protein leaves the lamellae physically intact, the possible involvement of cuticular protein in the primary orientation of the chitin is not of course excluded. The combined thickness of one pair of daily growth layers is of the order of 10 p, varying with location in the skeleton. In favourable conditions, adult desert locusts deposit solid endocuticle day and night for 2 to 3 weeks after the final moult. When they are reared in control conditions (12 h of light, 75 to 150 ft candles, at 35", alternating with 12 h of dark at 25"), the night layers themselves each contain several lamellae, whereas the day layers are non-lamellate (Fig. 15).

FIG.16. Diagram of the chitin architecture of a locust hind tibia. The outermost layer, in black, is the exocuticle (covered by a very thin epicuticle); the innermost layer is the epithelium of epidermal cells. Between these two lies the endocuticle, in which the directions of maximum and minimum refractive indices for the various chitin fibril systems seen in various sections are indicated by biaxial indicatrices (drawn t o the same line thickness as the relevant fibril system). 'TS: transverse section; LS: longitudinal section; OTS: oblique transverse section (from Neville, 1965b).

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A . C . NEVILLE

Chitin is present throughout the endocuticle in three components (Neville, 1965b) (Fig. 16): (1) lamellae in each night layer. Chitin oriented parallel to the cuticle surface; (2) longitudinal fibrils running along the leg or wing vein axis. These fibrils comprise the non-lamellate day layers, and when seen in transverse section (looking along the molecular chains) they are isotropic; (3) radially arranged pore canal filaments which complete the threedimensional chitin framework.

2. The chitin orientations in a locust hind tibia reared in constant day conditions The structural organization of locust cuticle can be controlled by varying environmental conditions at the time of its deposition (Neville, 1963d, 1965a, b). If locusts are grown in constant day conditions (constant light, 75 to 150 ft candles at 35'7, chitin lamellogenesis is inhibited, and the result is a non-lamellate endocuticle. Once deposited, neither lamellate nor non-lamellate endocuticle changes (with regard to the presence or absence of lamellae). The structures described cannot be artifacts since they are also seen in hand sections of living material. Any combination of lamellate and non-lamellate layers can be produced by altering conditions appropriately. Chitosan-iodide staining, which was also confirmed by chromatography, shows that chitin is by no means restricted to lamellate layers in endocuticle. Equally aged samples of control-reared (alternately lamellate and non-lamellate) and constant day reared (non-lamellate) locust hind tibiae were weighed before and after deproteinization and found to contain the same quantity of chitin (28.7% and 28.0% respectively). Thus chitin lamellogenesis is mainly a periodic process of permanent change in orientation of chitin crystallites occurring during deposition. Chitin lamellogenesis in locust cuticle is independent of chitin synthesis. In constant day reared non-lamellate endocuticle the chitin, which would have been oriented as lamellae during the nights, is now oriented as extra fibrils along the leg or wing vein axes. It could be argued that lamellae too closely spaced for resolution in the light microscope might be present in constant day endocuticle. However, the lamellae of exocuticle are too closely spaced for resolution but they do of course cause the exocuticle to be strongly birefringent parallel to its surface. Constant

CHITIN ORIENTATIONI N CUTICLE AND ITS CONTROL

239

day endocuticle has a low birefringence parallel to its surface, confirming the absence of lamellae. This has also been confirmed in the electron microscope. 3. Involvement of a biological clock Daily growth layers are still produced in the solid endocuticle even when locusts are reared at constant temperature in permanent darkness, indicating that the process is governed by a biological clock whose mechanism lies within the body of the insect. In darkness at 36" this clock persists for at least two weeks with a free-running period of about 23 h, so that it gains 1 h per day upon the astronomical clock: after 12 days it is exactly out of phase with it (Neville, 1965d). The locust clock is not therefore triggered by an environmental event occurring at the same phase point on each day and is, thus, circadian. Further evidence for this is shown by the almost negligible effect upon the free-running period of separate experiments performed at constant temperatures between 22" and 38". Temperature independence is a universal property of circadian clocks. The Qlo (temperature coefficient) between 26" and 36" of the clock which times chitin lamellogenesis in Schistocerca is 1.04 (calculated by using the reciprocal of the period to indicate the rate at a given temperature). This agrees well with Qlo values for circadian clocks in other organisms including insects (Sweeney and Hastings, 1960). The circadian rhythm of chitin lamellogenesis has been found to persist in cave crickets (Dolichopoda linderi) from the PyrCnCes living in an almost constant habitat of darkness at 13" (Neville, 1965d). It has proved possible to study the effects of various specific environmental conditions which uncouple the clock from the orientation mechanism in various cells. The experimental insects conveniently record the results in their cuticular deposits for later analysis. The fact that some epidermal cells uncouple more slowly than others at various light intensities (see below) shows that this is in fact an uncoupling process rather than an actual stopping of the clock. Chitin lamellogenesis in the cockroach Periplaneta americana and chitin crossed fibrillar orientation in the toe-biter water bug Hydrocyrius colombiae are both governed by circadian clocks (Neville, 1965d), but by contrast with the locusts Schistocerca and Locusta, the daily organization could not be uncoupled from the clock either by constant day or constant night conditions. Factors involved in uncoupling were therefore further investigated in locusts.

240

A. C . N E V I L L E

I thickened non-spine side

I cn

B

'

spine side cells secreting thickened cuticle o f

mm non-spine -bear/ng side o f ribla

ims cells secreting spine -bearing Tide o f tibia

A

FIG. 17. A. Diagram of a locust hind tibia t o show the locality of the proximal region mentioned in the experiments (divisions 0 t o 2). B. An outline of a transverse sectioa through the proximal region of the hind tibia shown in A. Exocuticle is black: stippled endocuticle is that in which continuous lamellation can be produced by growth in lob temperature at constant illumination. The non-stippled endocuticle is that in which the absence of lamelfation can be produced by growth in high light intensity at constant temperature (from Neville, 1965d).

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4. Uncoupling chitin lamellogenesis from the clock Locust epidermal cells differ in their response to specific imposed environmental conditions according to their location in the integument (Table V). In those cells secreting the thickened proximal region of the hind tibia (Fig. 17) constant low (for locusts) temperature (22-25") uncouples chitin lamellogenesis from the circadian clock provided that illumination (light or dark) is constant also: the result is continuously lamellate endocuticle (Fig. 18). In the cells producing the rest of the hind tibia, constant light acts as an uncoupling factor, provided that temperature (high or low) is constant also: the result in this case is continuously non-lamellate endocuticle (Fig. 26). These results on Schistocerca have been confirmed in Locusta migratoria. Hence with a combination of constant low temperature (22") and constant high light intensity (75 to 150 ft candles), the endocuticle of the thickened proximal region of the hind tibia is continuously lamellate, whilst the endocuticle of the rest of the hind tibia of the same animal is non-lamellate. If the response were in one cell, one would suspect a bistable state which could be triggered in ei her direction (lamellate or non-lamellate) and which was normally c upled to the circadian clock mechanism (so producing lamellate and non-lamellate endocuticle in alternating layers). However, this bistability cannot be found in any one given cell, but rather each epidermal cell may be biased in one particular direction. Thus, the results in Table V show that the cells secreting the locust radius plus media wing vein are uncoupled (towards non-lamellation) from the clock by any combination of constant conditions (e.g. cold night, cold day, warm night, warm day). The cells producing the

i

FIG. 18. Diagram of a transverse section of locust hind tibia1 cuticle showing the effect of continuous low temperature upon the proximal region shown in Fig. 17. Exocuticle is represented by closely spaced lamellae: endocuticle by lamellae drawn further apart. There is a switch OHof lamellogenesis at emergence.

TABLEV Analysis of environmental factors uncoupling chitin lamellogenesis from the circadian clock in various locust cells Cell location

Thickened proximal region of hind tibia Rest of hind tibia

Radius plus media wing vein

Control rhythm 12 h at36"Cand 75 to 150 ft candles a!ternating with 12 h at 26" C and dark Circadially lamellate and nonlamellate Circadially lamellate and nonlamellate

Circadially lamellate and nonlamellate Cuticle beContinuous tween radius lamellation plus media and cubital wing veins

(a) Combinations of Constant Constant cold day cold night 26" C and 26" C and 75 to 150 dark ft candles

constant conditions Constant Constant warm day warm night 36" C and 36" C and 75 to 150 dark ft candles

Continuous Continuous Circadially lamellation lamellation lamellate and nonlamellate NonCircadially Nonlamellate lamellate lamellate and nonlamellate

Circadially lamellate and nonlamellate Circadially lamellate and nonlamellate

Nonlamellate

Nonlamellate

Nonlamellate

Nonlamellate

Continuous Continuous Continuous Continuous lamellation lamellation lamellation lamellation

h,

G

(b) Abnormal rhythms 12 h at 12 h at 24 h at 36" C 75 to 150 36" C and alternating ft 75 to 150 ft with 12 h candles at 26" C. alternating candles alternating Constant with 12 h with 24 h light 75 to dark. at 26" C 150 ft Constant candles 36°C and dark Circadially lamellate and nonlamellate Alternately lamellate and nonlamellate but half as many layers as control

Circadially lamellate and nonlamella& Circadially lamellate and nonlamellate

Circadially lamellate and nonlamellate Circadially lamellate and nonlamellate

?

zm 5 m

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endocuticle between the radius plus media and cubital wing veins are permanently uncoupled from the clock (towards continuous lamellation) even in control conditions. Thus, the response to the interaction between environmental factors and the circadian clock is finally determined in the individual epidermal cell. The experiments leading to uncoupling from the clock so as to produce continuously lamellate endocuticle show that it is the process of timing of chitin lamellogenesis which has been uncoupled, rather than lamellogenesis itself having been inhibited.

5. Facultative and obligatory coupling Under constant day conditions there is an inhibition of chitin lamellogenesis in many, but not all, regions of the skeleton during cuticle deposition in the locust. The exceptions are always constant in location and may indicate regions where lamellae are functionally indispensable (Neville, 1965b). (1) In the rubber-like cuticle of the prealar arm and wing hinge ligaments, where their function may be to constrain forces in directions parallel to the cuticle (Weis-Fogh, 1960; Jensen and Weis-Fogh, 1962). (2) In the median ocellus and compound eye cuticle. (3) In the thickened region at the ends of the tibiae (to resist the twisting component during a jump?). (4) Along the edges of wing veins, where they may serve to strengthen the wing when it is twisted during flight. ( 5 ) In the innermost layer of the endocuticle. It has been suggested (Neville, 1965b) that this layer may be homologous with the noncalcified membranous layer of crustacean endocuticle (Travis, 1963). In the locust, this layer may thus be distinguished morphogenetically as well as morphologically. (6) In the exocuticle as a whole. This means that the same environmental conditions which uncouple chitin lamellogenesis from the circadian clock in the post-ecdysial skeleton (endocuticle), do not affect pre-ecdysial lamellation. Since pre- and post-ecdysial cuticles are made by the same single layer of cells, there may be a change at ecdysis in their susceptibility to changes in the external environment or to messages reflecting them. The cells behave as if they were obligatorily coupled to lamellation before emergence, then facultatively coupled to it via a circadian clock for the next two weeks of post-ecdysial deposits, finally becoming obligatorily coupled again during the secretion of the

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innermost layer of the endocuticle. Moult and intermoult morphogenetical events may be under different control systems. The possible function at the time of expansion of chitin lamellae in exocuticle has been mentioned in Section I11 D. 6. Temperature eflects The temperature coefficient (Qlo), between 26" and 36", for the circadian clock governing the timing of chitin lamellogenesis in locusts is 1-04 (Neville, 1965d). The Qlo over the same temperature range for increase in endocuticular thickness is approximately 2.0 (Fig. 19).

50

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FIG.19. The effect of temperature on growth of locust endocuticle measured from daily growth layers in hind tibiae at a well-defined locus.

Growth is rhythmical in arthropod skeletons and so is the organization of structure. Thus, in view of the involvement of circadian clocks in homeostatic mechanisms (Harker, 1964) it might be thought that if chitin morphogenesis is coupled to a temperature-compensated clock, the actual control of rhythmical skeletal structures would then become more independent of the environment. However, consideration of the discrepancy between the temperature coefficients for the timing of chitin lamellogenesis and for the rate of cuticle deposition, shows the homeostatic implication to be paradoxical (Fig. 20). In fact if the clock were really dependent upon temperature with a Qlo of 2.0, the mechanism would clearly be more effective as a homeostatic control of structure.

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FIG.20. Diagrams of transverse sections of locust cuticle to show the non-homeostatic influence of a circadian clock with a Qlo of 1.0 upon structure. (a) 48 h growth at To. (b) 48 h growth at T+ 10"in a system such as a locust with Qlo for growth of 2.0, and Qlo for circadian clock o f 1.0. (c) 48 h growth at T+10" in a homeostatic but hypothetical system with Ql0 for growth of 2.0 and Qlo for clock o f 2.0.

7. Determining the age of an insect

It is possible to use a known sequence of environmental conditions to produce and analyse modified cuticular structure, or conversely, to use cuticle structure as a method for determining the age of an insect in which daily cuticular growth layers have been established. For studies of population age dynamics, suitably large samples must be analysed and so a suitably rapid method was developed. This consists of hand-sectioning a hind tibia with a fresh razor blade under a binocular microscope, and counting the daily growth layers as seen with a microscope between crossed polaroids. Such an age determination takes about one minute to perform and has been used in the field on a population of nymphal and adult grasshoppers (Decticus verrucivorus) (Neville, 1963~).The results showed that the technique could be used satisfactorily by field workers. There is an alternative method of finding the age of a grasshopper which does not involve sectioning, and which can be done with the aid of an ordinary binocular microscope (A. C. Neville, unpublished). The apodemes on which insert the thoracic dorsal longitudinal muscles (prephragma, middle phragma and postphragma) grow in area by daily increments, so as to produce a daily banding (Fig. 21). The dorsal longitudinal muscles and their apodemes of insertinon thus grow in parallel fashion throughout the first 3 weeks of the adult instar. The system would be a good one in which to study the origin of muscle

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I

lrnm

1

FIG.21. Middle phragma of adult pterothoracic dorsal longitudinal muscles from locusts killed at emergence, 2 days after emergence, and 7 days after emergence. e marks the emergence line, and the figures denote postimaginal daily growth layers.

insertions into cuticle. The nature of the daily growth layers of phragmata has not yet been investigated. B . METABOLIC OSCILLATORS A N D “SWITCHES”

That structural periodicities occur in cuticles is a reflection of morphogenetical oscillators. The daily rhythm is not the only one involved in the deposition of locust cuticle. There are also the shorter period cycles of chitin lamellogenesis, occuring in both rubber-like and solid cuticles. 1. The rhythm of chitin lamella deposition in the rubber-like cuticle of the locust prealar arm The location of the prealar arm in locusts is described by Weis-Fogh (1960). This structure consists of chitin lamellae glued together and separated by layers of resilin. At 11 days after adult emergence, a prealar arm ligament contains 24% chitin by weight, consisting of about 130 discrete chitin lamellae (Neville, 1963b). Because the only other significant component present is resilin (76%), the prealar arm provides a good location to study events associated with lamellar deposition. The chitin lamellae behave chemically like chitosan after treatment with boiling KOH (Weis-Fogh, 1960), and electron micrographs (Elliott et al., 1965) reveal that each lamella exists as a discrete sheet. By making use of the daily fluorescent growth layers in the resilin of prealar arm sections, it is possible to analyse the stratigraphical sequence of chitin lamellae (Fig. 22). If the total number of chitin lamellae at given ages (as determined from pairs of photographs, as in

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FIG.22. Photographs, enlarged to the same magnification, of a strip from a locust prealar arm section viewed both in ultra-violet light to emphasize the daily growth layers in the resilin (left), and between crossed polaroids to emphasize the chitin lamellae (right). The daily growth layers are used to count the number of chitin lamellae present a t given ages of deposition. Time is indicated in days preceding emergence (negative) and days after emergence (positive), zero representing the time of adult emergence (from Neville, 1963b).

Fig. 22) is plotted against the dry weight of chitin present at the same ages (from Fig. 23), the result is the curve shown in Fig. 24. This shows that before and up to adult emergence, successivcly formed chitin lamellae are increasingly heavy; this i s associated with the initially rapid increase in length of the prealar arm throughout the preimaginal 10-kA.I.P.

4

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Age of odult (days) ( bl

(cl

FIG.23. Maturation growth of locust adult rubber-like cuticle. All points represent the mean of ten ligaments. The time of adult emergence is indicated by a broken line. The curves are extrapolated back t o zero weight a t - 3 days (before emergence) on the basis of preimaginal growth layer evidence (Neville, 1963a). (a) Growth curves for the increase with age in total dry weight of mesothoracic wing hinge and prealar arm ligaments. (b) Growth curves for the dry weight increase in protein (resilin) and chitin components of mesothoracic prealar a r m ligaments. (c) Growth curved for the dry weight increase in resilin and chitin plus fibrous protein, for the mesothoracic wing hinge ligament. The graphs show that only resilin is deposited between one week after emergence and full cuticular maturity (from Neville, 1963b).

period, which involves a large increase in area of successive lamellae. After emergence, the area of successive lamellae remains almost constant. The linearity of the curve in Fig. 24 after emergence indicates that the weight of all postimaginal chitin lamellae is constant at 0.123 pg. Each postimaginal chitin lamella thus constitutes a quanta1 unit. The frequency of deposition of chitin lamellae can be estimated from pairs of photographs like those in Fig. 22, and follows an age function as plotted in Fig. 25. The period between formation of two successive lamellae is initially less than 1 h, increasing to 8 h at 11 days after emergence.

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2. The rhythm of chitin lamellogenesis in solid endocuticle Whereas the frequency of chitin lamella formation in locust rubberlike cuticle could be determined by a metabolic oscillator, which operates by switching chitin synthesis on and off as it switches resilin synthesis

FIG.24. The total number of chitin lamellae at given ages (indicated by figures on the curve) as a function of weight of chitin present in a locust prealar arm ligament at the same ages. The number of lamellae present with respect t o age was determined from pairs of photographs like those in Fig. 22, and the chitin weights were obtained from Fig. 23. The linearity of the curve after emergence shows that each chitin lamella weighs the same, in this case 0 1 2 3 pg. Age in days negative before adult emergence (broken line), positive after emergence (from Neville, 1963b).

FIG.25. The frequency of chitin lamella deposition in the adult locust prealar arm ligament, expressed as number of chitin lamellae deposited in 24 h periods with respect to age. Vertical broken line again denotes emergence (frcm Neville, 1963b).

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off and on, the situation in locust solid endocuticle is different. Even when chitin lamellogenesis is prevented by constant day uncoupling from the clock, chitin synthesis clearly continues, giving rise to pore canal filaments and longitudinal fibrils along leg or wing vein axes. Here the oscillator controlling the frequency of chitin lamellogenesis operates upon the primary chitin orientation process, and not upon chitin synthesis. The time required to determine the organization of a single chitin lamella was measured experimentally (Neville, 1965b) as follows. Locusts were reared in constant day conditions (35" plus 75 to 150 ft candles) for a week following adult emergence, so that they produced a non-lamellate tibial endocuticle. A short spell of night

FIG.26. A sample of non-lamellate locust tibial endocuticle in transverse section, produced by rearing the locust in constant day conditions for a week following emergence, broken only by 15 min night in the middle of the wcck.

FIG.27. As for Fig. 26 but with 30 rnin of night conditions allowed in the middle of the week of constant day. The organization of a single lamella is determined during the 30 min night. Scales 50 p (both from Neville, 1965b).

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conditions (25" plus dark) was allowed in the middle of this period (midnight at the end of the fourth experimental day). A 15 min spell of night produced no lamellae, the resulting endocuticle being nonlamellate throughout (Fig. 26). However, with 30 min, a single lamella was organized (Fig. 27). Hence 15 to 30 min are required to determine the organization of a locust endocuticular chitin lamella. Condoulis and Locke (1966) have counted the number of chitin lamellae present at different stages of deposition in electron micrographs of the cuticle of fifth instar larvae of the wax moth, CaZpodes ethluis. The larvae were kept in constant darkness at 22" throughout the fifth TABLE V1 Number of chitin lamellae present at various stages of deposition of the cuticle in fifth instar larvae of the wax moth (Calpodes erhlrtis). From Condoulis and Locke (1966) Age after ecdysis to fifth instar (h)

Number of lamellae present

25 52 73 100 139 142

25 44 130 288 307 420

TABLE VII Average times for formation of lamellae at various stages of deposition of fifth instar larval cuticle of the wax moth (Calpodes ethluis), calculated from Condoulis and Locke (1 966) Period (h) 0-25 25-52 52-73 73-100 100-1 39 100-142

Number of lamellae

Average time/lamella (min)

25 19 86 158 19 132

60 85 15 10 12319

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instar. The results of this investigation are presented in Table VI. These authors conclude that the frequency of deposition of lamellae varies throughout the fifth instar. These results were obtained from different individuals killed at various stages, rather than by the counting of a chronological sequence in one animal as can be done in locusts. The average times of formation of lamellae for various 24 h periods are calculated in Table VII. The results are in fair agreement with the time of 15 to 30 min determined experimentally in locusts. In the antenna1 cuticle of the honey bee, Apis mellifera, Richards (1952) found 15 to 20 changes in the direction of chitin deposition in a single day (Fig. 5). This system must also involve a metabolic oscillator with a period lying maximally between 70 and 95 min. In all four cases (locust rubber-like cuticle, locust solid endocuticle, wax moth cuticle and honey bee cuticle) these short period metabolic oscillators are endogenous, since they all persist in constant conditions.

FIG.28. The pause in deposition of chitin lamellae shortly following adult emergence. seen in a section of a locust prealar arm ligament photographed between crossed polaroids. The chitin lamellae are shown in the extinct position against the swollen resilin, which shows strain birefringence (from Neville, 1963b).

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3. Metabolic “switches” Sections of the locust prealar arm (Fig. 28) show that there is a pause in chitin lamella formation during adult emergence, whereas the deposition of resilin continues (Neville, 1963b). Measurement of the chitin lamellar frequency before and after emergence shows that two lamellae have been missed, corresponding to a pause in chitin synthesis of at least 6 h. There appears, therefore, to be a metabolic “switch” controlling chitin synthesis. A similar switch operates in the cells secreting another of the ligaments of the locust flight system, the wing hinge. The layered deposit can again be analysed stratigraphically, making use of daily fluorescent growth layers in the resilin (Neville, 1963b). Figure 29 shows that nothing but pure resilin is deposited in the ligament between one week after emergence and full adult maturity, deposition of chitin and tough fibrous protein having then ceased. This is confirmed by evidence from growth

chitin larnellae Pure resilin Tough fibrous protein with some chitin lomellce

I

0.5 mm

FIG. 29. The architecture of the locust wing hinge ligament. A transverse section of a mesothoracic wing hinge ligament from a locust 2 weeks old from emergence, with the various regions of material symbolized. The amounts of material already deposited at emergence, 1 and 2 weeks old are enclosed by the broken lines as shown. Camera lucida drawing from phase contrast and fluorescence microscopes (from Neville, 1963b).

2 54

A . C. N E V I L L E

curves (Fig, 23). Figure 29 shows that the end of chitin deposition spreads as a wave over the cells from one day before emergence until seven days after. A switch influencing chitin orientation at adult emergence in locust solid endocuticle is revealed by rearing locusts at low temperature (26") in the dark as described in Section IV A above. In the thickened proximal region of the hind tibia lamellation is then continuous (Fig. 18). At emergence there is a pause in chitin lamellogenesis, between the finely lamellate preimaginal exocuticle and the more widely spaced lamellae of the postimaginal endocuticle. At 26" the non-lamellate region is 5 p wide, representing six postimaginal lamellae missing. These results from locusts suggest the involvement of a metabolic timing device at emergence which causes: (1) chitin synthesis to turn off and on again a few hours later in the cells producing the rubber-like cuticle and (2) the chitin orientation mechanism to be affected similarly in the cells producing the solid endocuticle. c. " D E R M A L " L I G H T S E N S E When locust adults are reared in constant day conditions the endocuticle over most of the hind tibia is non-lamellate. When they are reared in constant warm night conditions the same parts of the endocuticle are circadially organized with lamellate zones alternating with non-lamellate ones. Therefore, since the only difference between these conditions is illumination intensity (temperature being constant at 36"), there should be a light threshold at which uncoupling of chitin lamellogenesis from the circadian clock occurs. This was investigated by rearing freshly emerged locust adults in constant illumination at various intensities. The results (Fig. 30) showed that the lower the light intensity, the greater the number of days required to uncouple chitin lamellogenesis from the circadian clock. Even with 75 to 150 ft candles illumination, lamellae were still formed on the first astronomical night of the experiment, but not on the second night. These results suggest the possibility of the accumulation of a hormone or other metabolite which is regulated by light intensity. This is analysed further in Fig. 31, in which the number of chitin lamellae produced per night zone at a well-defined locus on the hind tibia is plotted for the various light intensities. The result shows that the higher the light intensity, the less the number of lamellae in each successive layer, until they disappear altogether. The above results were all obtained starting with freshly emerged adult locusts. However, the same number of days to switch off at a

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given light intensity is taken by adults at any age after emergence during the deposition period. Light intensity still determines the number of days of constant day conditions required for uncoupling even when the two compound eyes and the three simple ocelli have all been cauterized. The locusts might -0

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FIG.30. Graph showing the number of 24 h periods of constant day conditions at various light intensities which are required to uncouple chitin lamellogenesis from the locust circadian clock; measured at a constant locus on hind tibiae. Note the break in the abscissa between 10 and 74 ft cafidles.

therefore possess a “dermal” light sense which is capable of measuring light intensity. Dermal light senses are classically regarded as triggering short term behavioural effects (Steven. 1963). This long term metabolic effect in locusts is, therefore, more like photoperiodic responses. Preliminary results for an equal energy action spectrum for the process also give results resembling typical photoperiodic action spectra in insects and mites (e.g. Lees, 1953). The pigment responsible for light absorption absorbs between 4350 A and 5200 A. This tallies with the absorption spectrum for /%carotene which is abundant in locusts, although this correlation could be entirely fortuit6us. However, it is pertinent to note that while carotenoid metabolism is unaffected by 10*

256 A. C . NEVILLE temperature in locusts, high rearing temperatures cause locust adults to appear more yellow in colour @carotene) because less ommochromes are then present to mask them (Goodwin, 1952). Conversely, adult locusts reared at low temperatures have more ommochromes I

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FIG.31. Graphs showing the number of chitin lamellae produced on each successive night a t a defined locus in hind tibiae of desert locusts reared in various light intensities a t various temperatures.

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which mask the colour of the 8-carotene. Experiments in which locusts were reared in various combinations of constant temperature and light intensity might be revealing, if analysed in terms of the length of the uncoupling period in specific cells. An independent approach could be made by working out a detailed action spectrum curve for the process. A specific test for the involvement of /3-carotene would be to rear the locusts on a 8-carotene-free diet in constant day conditions. The location of the site of light reception in this uncoupling reaction is not yet known for certain. Locusts in which the dorsum or whole of the head capsule was painted black still responded to light intensity. These very preliminary findings might indicate that the site of light reception is not the neurosecretory cells in the brain, such as have been implicated in photoperiodic responses controlling polymorphism in aphids (Lees, 1964). The recent demonstration of photosensitivity in the terminal abdominal ganglion of Periplaneta (Ball, 1965) suggests alternative lines of investigation. The possibility that the light intensity was actually being measured by the living green wheat which was fed to the locusts, which then consumed some specific metabolite in their food, was eliminated by omitting green food from the diet. When fed only on moist wheat bran, newly emerged adults were found to be able to deposit endocuticle for 3 days on food reserves. When they were allowed to deposit on reserves in constant day conditions (35" plus 75 to 150 ft candles), uncoupling of chitin lamellogenesis from the clock took place after one night, as usual. To show that it was not simply starvation of green food which caused this uncoupling, further freshly emerged adults were reared in control conditions (alternating hot days and cold nights) also without green food. Individuals treated in this way produced three lamellate and three non-lamellate layers of endocuticle in three days as expected. It may safely be concluded that: (1) locusts measure light intensity over long periods by some agency other than with their compound eyes and ocelli; (2) the food plant is not concerned in this response; and (3) the measured light intensity between wavelengths of 4350 and 5200 A causes a long term graded uncoupling of the orientation of chitin into lamellae from the circadian clock which normally controls its timing. D . IMPLANTATION EXPERIMENTS

A series of experiments was performed upon cylinders of hind tibiae cut from living individuals and implanted loose into the haemocoel through a cut in the intersegmental membrane between the second and

FIG. 32. Transverse section of a locusl hind tibia photographed between crossed polaroids. The leg was implanted into the haemocoel two nights after adult emergence. exo = exocuticle; pre = endocuticle grown before implantation; r = random endocuticle grown during reorientation of epidermal cells; post = four nights of endocuticle grown during implantation pcriod, after reorientation of cells.

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third abdominal sternites. The wounds were sealed with wax and one set of locusts (3 days after emergence) placed into control conditions for 7 days following the surgery. Subsequent dissection and sectioning showed that the implants had taken and that endocuticle had been deposited subsequent to implantation. A zone of badly oriented material separated the pre- and postimplant deposits (Fig. 32). The post-implant growth was lamellate and non-lamellate in alternation but in some cases the non-lamellate layers were birefringent in transverse sections (TS) of the leg (Fig. 32). This means that the longitudinal chitin fibrils, normally isotropic in TS, were not perfectly oriented along the axis of the implanted cylinder. Chitin pore canal filaments ran radially through the post-implant deposits as usual. The implanted epidermis divides and migrates so as to encapsulate the whole of the original implanted cylinder. During migration the deposits are poorly oriented on both sides of the original cylinder. Short cylinders (about 2 to 3 mm long) are most effective since the cells can then join up fairly quickly. Postimplant endocuticle in cylinders of this size showed three lamellate layers less than the number of days of post-implant deposition, indicating that the migration and encapsulation took 3 days, and also that the subsequent lamellate and non-lamellate layers on both inside and outside of the original cylinder are still organized in a daily manner. As in controls, each lamellate layer contains several lamellae. Repeating the experiments with locusts in constant warm night conditions (dark at 35") revealed that chitin orientation still shows a circadian rhythm even in implanted deposits. The unoperated hind tibia on each recipient animal here acted as a control. Reciprocal implantations were carried out between Schistocerca gregaria and Schistocerca paranamense, each member of a pair acting as both donor and recipient. The implants still organized chitin lamellae according to a circadian rhythm while growing in a different species. These experiments show that neither mechanical nor nervous connections to the epidermis are necessary for the orientation of chitin into lamellae or into parallel fibrils. The deposits which are organized during cell migration in implants are badly oriented. They give rise to a plane of cleavage between the pre- and post-implant deposits. This provides a means of separating endocuticle from exocuticle by dissection. Furthermore, after migration, the cells encapsulate the implanted cylinder with endocuticle which is again easily separable. The technique could be used for the production of quantities of pure endocuticle for physical and chemical studies, since these have so far been restricted to combinations of

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exocuticle and endocuticle. The production of a badly oriented weak zone of cleavage by migrating cells emphasizes the importance of the careful cellular control of orientation of cuticular components in the production of a strong skeleton. E. N E R V O U S C O N T R O L

The evidence from implantation experiments (Section 1V D) clearly eliminates the possibility that the timing of chitin lamellogenesis in locusts is under nervous control, since the implanted cylinders were completely denervated. Similarly, it rules out the possibility of orientation control by the passage of neurosecretory material along axons to the epidermal cells in locusts. Maddrell (1962) and Nunez (1963) demonstrated the involvement of the nervous system in controlling the increase in size during feeding in Rhodnius fifth instar larvae. Cutting the nerves to one side of the abdomen reduced distension during feeding on that side. It also produced corresponding differences in the orientation of chitin micelles between the two sides which Nunez attributed to a decrease in cohesion between chitin crystallites due to plasticizing changes in the structural protein. Bennett-Clark (1961) and Maddrell (1966) consider this to result from a change in cuticle hydration. S. H. P. Maddrell (personal communication) has shown that this response may also be demonstrated at the segmental level and finds evidence for the release of neurosecretory granules at the ends of the relevant axons (Maddrell. 1965). The chitin reorientation is permanent and still occurs whether feeding immediately follows neurotomy or one month later. The involvement of muscles in this process is clearly ruled out since these are involuted except at the moult (Wigglesworth, 1956). The above experiments provide a good example of one type of secondary reorientation, with its own control system (see Section V A and B). F. DISCUSSION

The organization, composition and spatial arrangement of the chitin deposits of locust cuticle provide a reliable and detailed chronology of metabolic events affecting its deposition, and of their temporal control. At a higher level (weeks and months) temporal control is well known to affect insect growth and diapause by combinations of environmental photoperiodic changes with interval timers (Lees, 1960). It is now

CHITIN O R I E N T A T I O N I N CUTICLE A N D ITS CONTROL -- ---- - -- - ----- - 1 Constant light (at constant temperature)

261

/ /

I

/

/ 3

/ /

Phase

/ /

I

Basic chitin

I I I

mechanism

& orientation J

FIG.33. A possible scheme for the physiological control of chitin orientation in locusts.

Trehalose in blood

1 1

Glucose Phosphorylation

Glucose-6-phosphate

Fructose-6-phosphate ( ?) Glutamine Amination

IC

Glutamic acid

Glucosamine-6-phosphate Acetylation

lc:zl-CoA-Lipids

or TCA intermediates

~-Acetylglucosamine-6-phosphate

J.

N-Acetylglucosamine-1-phosphate ( ?) Activation by conjugation with

UDP Uridine diphosphate N-Acetylglucosamine

b-;-Acetylglucosamine),, Polymerization (chitin synthetase)

Chitin (~-Acetylglucosamine),,+ 1

FIG.34. Pathway for chitin synthesis (compiled from various sources cited in text).

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becoming apparent that the timing of metabolic events in insects is significant down to lower levels (days and minutes). Although such intervals are longer than the range of operation of most nervous mechanisms we should not restrict the search for the oscillators involved to hormones and release of metabolites, since long term nervous phenomena (Bruner and T a w , in press) and even circadian rhythms (Strumwasser, 1965) are now known in Aplysia neurons. In one case, that of chitin lamellogenesis in locust cuticle, direct evidence against neural timing has been demonstrated. A hypothetical working diagram of the interaction between internal and external factors involved in chitin orientation control in locusts is given in Fig. 33. This figure is modelled on the lines of the general scheme of Locke (1964; Fig. 34), and is intended to serve as a conceptual framework. It should be borne in mind that the events within the dotted rectangle might turn out to be intracellular, in which case the postulated hormone would be reduced to an intracellular metabolite. If, however, either the clock or the site of light reception, or both, are localized in specific regions of the body, then hormones integrating the responses to environmental changes would be a necessary corollary. V . O R I E N T A T I OM NE C H A N I S M S One source of confusion leading to the classical controversy as to whether protoplasmic or mechanistic forces are responsible for determining macromolecular orientations, may be clarified by distinguishirig between primary and secondary orientation mechanisms. By “primary orientation ” is implied the positioning of a macromolecule by intermolecular forces acting between itself and other macromolecules in an aqueous or liquid crystal environment, whose composition may be under cellular control. “ Secondary orientation” may then be thought of as a rheological re-orientation caused by some physiologically derived force (e.g. muscular force, mechanical stress, blood pressure, feeding expansion forces or plant cell turgor pressure). It is probably associated in several cases with plasticization of protein and proteinchitin complexes. Both primary and secondary orientation mechanisms may occur simultaneously, and should not be considered as alternatives. Studies of plant cell-wall deposition have been confused by the difficulties of distinguishing between deposition of microfibrils in specific controlled orientations and simultaneous reorientation caused by changes in cell shape. In this respect we can draw an analogy between primary plant

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cell walls and insect presumptive exocuticle, and also between secondary plant cell walls and insect endocuticle. Locust endocuticle is ideal for the study of primary orientation, since, unlike in Rhodnius, it is not subjected to expansion. The chitin orientations can be changed permanently, but only at the time of chitin deposition (i.e. during primary orientation). The morphogenetical studies on locust cuticle reviewed above, together with the results of isotopic labelling and autoradiographic analysis of Calpodes cuticle (Condoulis and Locke, 1966), show that chitin, once deposited, is permanent and not labile as is some of the endocuticular protein. The chitin in the cell walls of unicellular bristles of Drosophila (Lees and Picken, 1945) and of the unicellular scales of Ephestia (Picken, 1949) shows primary axial orientation from the time of their outgrowths, but this orientation is secondarily enhanced by plastic flow during the elongation of the cells. A. PRIMARY ORIENTATION

I. Evidence for extracellular polymerization In order to be able to determine where the primary orientation of chitin is carried out, it is necessary to know whether chitin polymerization is extracellular or intracellular. This involves some consideration of the pathway of chitin synthesis in arthropods. The state of present knowledge is summarized in Fig. 34, based upon Candy and Kilby (1962), Jaworski et al. (1963), Gilmour (1965), Carey (1965) and Lipke et al. (1965), Condoulis and Locke (1966), The synthesis pathway is very similar to that of cellulose in plants. The scheme of addition of activated monomers to a growing polymer chain is compatible with the possibility of extracellular polymerization. Extracellular polymerization of cellulose by the bacterium Acetobacter xylinum is well established (Colvin and Beer, 1960, Ben-Hayyim and Ohad, 1965). Extracellular chitin production has been demonstrated in fungi (Castle, 1945). The evidence for extracellular polymerization and crystallization of chitin in arthropods derives largely from specialized sources : (1) Picken et al. (1947) found that the laminated cocoon of the beetle Donacia, which is aquatic and spun from a viscous fluid, contained both chitin and sclerotized protein; (2) Rudall (quoted by Picken, 1960) showed that the “silk” spun from solution by pronymphs of Mantis contains chitin; (3) the formation of the peritrophic membrane from a semi-fluid secretion is well known (Vignon, 1901 ; Wigglesworth, 1930; Picken, 1960); (4) B. L. Gupta (personal communication) has shown that

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a parabolic chitin fibrillar system can be developed in Copepod spermatophores, after secretion as a fluid; (5) there is also good evidence that the cuticle in Podura aquatica (Collembola) is formed extracellularly, for Noble-Nesbitt ( 1963b) has produced electron micrographs which show that the plasma membrane of the epidermis remains intact throughout the complete moulting cycle; (6) Taylor and Richards (1965) interpret the strongly PAS positive reaction of the subcuticle next to the epidermal cells as being due to non-acetylated or short chained chitin redues. They comment that this is evidence for polymerization of chitin and for chitin-protein cross-linking in the subcuticular region. 2. The region of primary chitin orientation The question whether chitin organization is determined in the zone of deposition adjacent to the cells, or whether it is controlled by postsecretory chemicals and stresses operating at considerable distance from them (Richards, 195I ; Picken, 1940, I960), has been discussed by Neville (1965b). The literature contains several claims that cuticular lamellae may be formed at a distance from the epidermis, while separated from it by an appreciable thickness of intervening cuticle. Examples include: ( I ) the exocuticle of Sarcophagu larvae (Dennel, 1946); (2) Cicada eye lens cuticle (Verhoeff, 1926-32; Richards, 1951); (3) organization of balken fibres in Calandra exocuticle (Reuter, 1937). However, the appearance of interposed lamellae between pre-existing ones as evidenced by light microscopy, need not necessarily involve an increase in number of lamellae. Lamellae previously too close together to be individually resolved could become visible simply as a result of cuticular swelling. Condoulis and Locke (1 966) have demonstrated that the first-formed lamellae in the untanned endocuticle of the wax moth Calpodes later enlarge with growth by intussusception so as to lie slightly further apart. They correlate this with the diffuse incorporation of certain amino acids (proline, tyrosine, cystine, histidine and tryptophan), detected by isotopic labelling and autoradiography. They point out that the amount of intussusception is greater than it seems, since the cuticle expands three times in area with feeding throughout the fifth instar, and this must be compensated by an equivalent factor of intussusception just to maintain the original lamellar separation. The aquatic cocoon of the beetle Donacia is a specialized case, forming lamellae after being secreted as a fluid while separated from the epidermis by intervening cuticle (Picken et a/., 1947). These lamellae are, however, far less precisely organized than in cuticles (A. C. Neville, unpublished). In the exocuticle of the Collembolan Podura aquatica,

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265

lamellae are dis-organized at a distance from the cells prior to hardening, whilst the intervening endocuticular lamellae remain intact (NobleNesbitt, 1963b). Continuity of lamellae over large areas of body surface, and their production in a successively one-by-one manner has been established for locusts (Neville, 1965b) and for Podura (Noble-Nesbitt, 1963b). In both locust rubber-like cuticle (Neville, 1963b) and solid endocuticle (Neville, 1965b), the most recently formed chitin lamella in locusts killed at any stage of deposition, is birefringent and crystalline, lying adjacent to the epidermis. This evidence conflicts with the views of Richards and Anderson (1942) who regarded the cuticular lamellae of cockroaches as being formed subsequent to secretion by a chemical phenomenon, and not as representing an unbroken sequence of deposition. Condoulis and Locke (1966) found that tritiated glucose is incorporated into chitin in the endocuticle of waxmoth larval cuticle; subsequent autoradiographs show it occurring as sharply defined layers, which vary in thickness with the length of the incorporation period. When larvae were killed and autoradiographed shortly after injection, the labelled band of chitin occurred only at the interface between the cuticle and the epidermis. Electron micrographs also demonstrate that the organization of chitin lamellae occurs in close contact with the epidermal cells (Galleria larvae, Tenebrio pupae, Calpodes pupae; Locke, 1961). The most recently formed region of cuticle is about 1 to 2 p in thickness and is more coarsely granular and less well oriented than fully formed cuticle. The granules are seen to be in the process of orienting into parabolae as in fully formed lamellae. These results allowed a more plausible interpretation (Locke, 1961) of the function of the so-called subcuticular or Schmidt’s layer, which had been considered as a non-chitinous glue (Schmidt, 1956), as the most recently formed layer at any age (not yet fully organized). It seems reasonable to suppose that the chitin which is not yet stabilized by cross-linking and hydrogen bonding to protein and to itself, would be dispersed by treatment in boiling NaOH, so leading to an anomalously “negative” result in subsequent chitosaniodide tests. We may conclude that the region of primary orientation of chitin is in the zone of deposition adjacent to the cells. B . SECONDARY ORIENTATION

It is well established that if tension is applied t o a gel, anisodiametric crystallites rearrange so as to lie parallel with the applied tension. Thus

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the study of secondary orientation effects involves the field of plasticiza tion of cuticle. Secondary orientation increases the form birefringence parallel to the orienting force and this has been shown in artificially drawn arthropodin fibres (Richards and Pipa, 1958), artificial chitin fibres (Clark and Smith, 1936), precipitated actomyosin threads (Szent-Gyorgyi, 1941), plant cells stretched in their direction of growth (Bonner, 1935), stretched blowfly larval cuticle (Fraenkel and Rudall, 1940) and silkworm threads polymerized under tension (H. Mark, 1934, quoted in Houwink, 1958). Mark made fibres of extremely well oriented natural silk by forcing a silk worm to spin its thread under tension. This was done by means of a motor which wound up the thread while drawing it out of the larva. 1 . Anisotropic skeletal strains Are anisotropic skeletal strains involved in the orientation of chitin crystallites in insect cuticle as was suggested by Picken (1940) and Picken et al. (1947) ? Such mechanistic theories have often been invoked to explain the predominant orientations of cellulose crystallites in growing plant primary cell walls (Castle, 1936, 1937) and of chitin crystallites in the walls of fungi (van Iterson, 1937). However, Bonner (1 935) showed that when an Avena coleoptile parenchyma cell elongated by 100% during growth, no obvious orientation changes occurred, whereas a mechanical extension of only 10% caused marked reorientation. Conversely, in bean stems, extensive deposition of longitudinal microfibrils still occurred when elongation had been reduced experimentally (Setterfield and Bayley, 196 1). During cuticular deposition, the locust hind tibia is subjected to various forces acting through the skeleton as a result of posture and locomotory movements. It may be regarded as a cylinder subject to extension and compression so that the shearing forces act maximally in directions at 45" to its long axis. (In a solid plastic cylinder, diagonal cracks develop at 45" to the axis during compression tests.) If such forces were to play a part in orienting the chitin crystallites at the time of their deposition, the resulting orientations would also be expected to lie at 45" to the long axis. Form birefringence analysis (Neville, 1965b) and X-ray diffraction analysis (Rudall, 1963) of locust leg and wing vein cuticle show that this is not the case. This, taken together with the results of the implantation experiments reported above, suggests that such forces do not play an important part in chitin orientation in locusts.

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267

2. Muscular forces The formation of the puparium from the larval cuticle in Diptera involves muscular contraction (Pantel, 1898), together with dehydration, secondary reorientation and closer packing of the molecules (Fraenkel and Rudall, 1940, 1947). X-ray diffraction shows that the chitin chains (which in the larva are oriented parallel to the cuticle surface but otherwise at random) come to have a preferred orientation circumferentially around the diameter of the puparium. This orientation provides for easy cleavage when the adult emerges, by breaking hydrogen bonds and van der Waals’ forces perpendicularly to the chitin chains. Ligation of the larva prevents contraction of the posterior part of the puparium (Fraenkel and Rudall, 1940). McLintock (1964) provides additional evidence for the involvement of muscles in this reorientation ; he notes that in Hypoderma bovis contraction affects only the abdominal tergites, whereas in Cuterebra it affects only the sternites. The families Oestridae and Cuterebridae also show that reorientation is independent both of hardening and darkening, since the sequence follows the order darkening, contraction and hardening, In Musca autumnalis hardening follows reorientation, but no darkening occurs. Maloeuf (1935) suggested that tonic muscular contraction at the last moult was involved in the formation of the long, highly oriented apodemes of dragonflies. However, in view of the fact that dragonfly flight muscle has a very high parallel elasticity present at its naturally occurring length in the thorax (Weis-Fogh, 1959), it is suggested that the extension of the flight muscles which occurs when an adult dragonfly expands, would place a considerable passive elastic tension upon the as yet untanned cuticle of the apodemes, sufficient to cause reorientation of the chitin and protein. Such a method would also be more economical in terms of energy than would sustained tonic contraction. The involvement of certain muscles in apodeme formation of adult bees (Daly, 1965), may also involve parallel muscular elasticity. Investigation of this possibility seems desirable. 3. Expansion at the moult Kroon et al. (1952) compared X-ray diffraction pictures of unexpanded and expanded wings of the small tortoiseshell butterfly, Aglais urticae. They found that chitin orientation was random in the unexpanded wing, whereas after expansion it was oriented along the wing vein axes. They considered that the reorientation of the chitin was an essential factor in stiffening the wing veins. Cottrell(1964), however,

268

A . C . NEVILLE

produces arguments to show that sclerotization is the major factor contributing to cuticular hardness. The exocuticle of the adult locust hind tibia contains chitin lamellae which are too closely spaced for resolution by a light microscope, but detectable by deproteinization and pressure (Neville, 1965b) and in the electron microscope (A. C. Neville, unpublished). They give rise to the strong positive birefringence of the exocuticle parallel to its surface. These lamellae are absent from the middle zone of the thickened side of the proximal region of the hind tibia (Fig. 35). This zone consequently lacks birefringence parallel to the surface. The chitin pore canal filaments are bent during emergence in this non-lamellate region, but remain straight in the lamellate regions (Fig. 36). Pharate adults, sectioned just prior to emergence, also show the pore canal filaments

FIG.35. Diagrammatic cross-section of a locust hind tibia to show the location of the non-lamellate regions of the exocuticle. The insets show the bending of the pore canal chitin components.

running a straight course through the non-lamellate regions. Thus, the distortion (secondary reorientation) occurs during expansion, and the observations show that the chitin lamellae contribute towards the control of the reshaping which occurs during expansion, by stabilizing the regions where they occur. There is evidence that the epicuticle is one factor concerned in controlling the shape of expanding Rhodnius

FIG.36. Transverse section of a locust hind tibia photographed between crossed polaroids to show the bending of the chitin pore canal components in a non-lamellate region of exocuticle.

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cuticle (Bennett-Clark, 1963). Local prehardening before ecdysis (Cottrell, 1962), local differences in thickness and chitin orientations (Neville, 1965b) probably contribute significantly also. 4. Expansion during an intermoult Caterpillars grow within a stadium by the inflation of a plastic skeleton (Locke and Greenberg, 1963 and quoted in Locke, 1964). The cuticle expands by the same amount axially and circumferentially (Condoulis and Locke, 1966) by 170 to 180%. This would not be expected unless the cuticle is mechanically anisotropic, since the hoop stress in a cylinder under uniform internal pressure is twice the axial stress (Timoshenko, 1951). Hence it might be expected that the chitin in a caterpillar is preferentially oriented circumferentially. The effect of the expansion forces during feeding upon reorientation of chitin in Rhodnius is reviewed above, in Section IV E. The secondary orientation of chitin during the intermoult growth of tracheae of Rhodnius has been interpreted in terms of expansion and buckling of the cuticulin layer (Locke, 1958) and has been reviewed by Locke (1964). Being invaginations, the tracheae are not restricted by the existing cuticle, so that in this case a growth force can act as a reorientation force. In summary, it can be said that there are several ways of experimentally interfering with both the primary and secondary orientation of chitin : (1) PRIMARY. Uncoupling the primary orientation mechanism from the circadian clock in locusts by various environmental treatments during deposition. (2) SECONDARY. (a) Ligation of pupating dipterous larvae (Fraenkel and Rudall, 1940). (b) Nerve sectioning in Rhodnius before expansion during feeding (Nunez, 1963 ; Maddrell, 1962, 1966). Crippling emerging endopterygote adults. C. PROTEIN ORIENTATION

In 1950, Rudall focused attention on the chitin-protein complexes of insect cuticle and upon the mechanism of their orientation. The hypothesis in the following Section (V D) represents an attempt to explain how these two aspects may be interrelated, as has been suggested by Kent (1964). It has been known for many years (Fraenkel and Rudall, 1940) that the crystal lattices of chitin and “/?-arthropodins” are dimensionally compatible (Fig. 3). The question as to whether the

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protein and polysaccharide chains are arranged parallel or mutually at right angles has, however, long been a controversial point (Rudall, 1950; Richards and Pipa, 1958; Rudall, 1963). Evidence for oriented protein in insect cuticle is to be found in several papers (e.g. Fraenkel and Rudall, 1940, 1947; Picken and Lotmar, 1950; Rudall, 1963). We do not yet know whether the chitin orients the protein or vice versa, or, as is suggested below, whether the process is a mutual orientation effect regulated in the deposition zone by the cells. One might expect that environmental changes which acted at the time of deposition and altered the primary orientation of chitin molecules, may also have changed the orientation or even the structure of the associated protein. That experimental interference with the environment at the time of deposition can, in fact, permanently modify protein structure has been postulated in the case of resilin (Neville, 1963b, 1965~).One line of approach would be to measure, by X-ray diffraction, any orientation changes in the protein of non-lamellate or all-lamellate samples of locust endocuticle in which the chitin orientation has been changed by environmental disturbance of the circadian clock. In view of the evidence for cross-linking of protein to chitin via histidine and aspartic acid residues (Hackman, 1960), and of the dimensional compatibility of the two kinds of macromolecule, Rudall's (1950) suggestion that the anti-parallel arrangement of a-chitin may be determined by the antiparallel arrangement of the /3-protein chains seems plausible. In support of the alternative crossed-grid theory, Hackman (1959) has suggested that such an arrangement would, when tanned by quinones, give a structure which would break but which would resist splitting. He notes that Lees and Picken (1945) observed that the sclerotized bristles of Drosophilu melunogaster break irregularly before diaphanol treatment, whereas after it the chitin which remains splits easily into fibrils. Tf we implicate protein orientation in the orientation of chitin, we should expect to find that the chitin chains associated with resilin in rubber-like cuticle would be randomly oriented (except for lying parallel to the epidermis), since resilin is a random three-dimensional macromolecular network of polypeptide chains. The chitin lamellae of rubber-like cuticle are in fact of this nature (Elliott et ul., 1965). The evenly spaced resilin cross-links (average separation 28 A; Andersen and Weis-Fogh, 1964), which are formed continuously with deposition (Neville, 1963b), present a network which cannot be penetrated by the forming chitin macromolecules. Secretion on to a pre-existing surface could account for the orientation parallel to the epidermis and in no other preferred direction.

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Another example of randomly oriented protein, in this case 8arthropodin, occurs in the oothecae of Blatta, which reveal a typical powder diagram by X-ray diffraction (Pryor, 1940). In connection with the concept of mutual orientation between chitin and protein it is interesting that this structure lacks chitin. D. H Y P O T H E S I S

The orientation hypothesis presented here is based upon a postulated cellular regulation of ionic concentrations in the deposition zone, which would affect orientation due to changes in charge interaction between the chitin and protein chains. It is speculative, but may serve as a working hypothesis since it is open to experimental testing. It has already been shown above that several mechanisms can be eliminated as possible candidates for control of primary orientation of chitin in insect cuticle, whilst remembering that they may well contribute to secondary orientation (e.g. skeletal forces, muscular forces, passage of nervous messages or neurosecretory material along axons to the cells). The evidence given above for the involvement of a circadian clock in primary orientation does not of course solve the problem of the ultimate orientation mechanism ; it merely indicates a higher order of control over it. In order to be tenable, any hypothesis on the ultimate mechanism for control of chitin orientation in cuticle must comply with the known facts established in Section IV. The mechanism must be capable of rhythmical control from the epidermal cells, both on a daily frequency (daily growth layers) and at higher frequencies (lamellae). It must also take into account the simultaneous formation of a chitin lamella over large areas of the epidermal surface with orientation occurring in the zone of deposition next to the cells. The cross-linking of polypeptide chains in resilin by amino acids (dityrosine and trityrosine; Andersen, 1964) is performed continuously in parallel with deposition (Neville, 1963b ; Fig. 37). This makes possible the precise cellular control of the regular arrangement of the stable covalent cross bridges and the lack of physical entanglement of the chains, which together account for the unique mechanical properties of resilin (Weis-Fogh, 1963; Andersen and Weis-Fogh, 1964). Such a method of cross-linking contrasts sharply with the more random crosslinking of man-made rubbers, in which the bulk process of vulcanization takes place after all the component rubber chains have been made. The classical tanning process of ordinary insect exocuticle is also a bulk-

C H I T I N O R I E N T A T I O N I N C U I I C L E A N D ITS C O N T R O L

-, ___ 0.2

0

273

I

1.0

rng resiiin (from samples of known age) FIG.37. Diagram establishing that the macromolecular network of resilin is crosslinked continuously as it is formed. Optical density measured at 315 m p of resilin ligament hydrolysates at different ages. Ordinate logarithmic with respect to fraction of light transmitted and linear with respect to concentration of the two fluorescent amino acids, dityrosine and trityrosine, which have absorption peaks near this wavelength. Abscissa, actual weight of resilin in samples of ligaments at known ages (ten per sample). 0 = prealar arm ligament: = wing hinge ligament; 1 = young prealar arms; 2 = young wing hinges; 3 = fully grown prealar arms; 4 = fully grown wing hinges (from Neville, 1963b).

+

tanning procedure. The occurrence of continuously controlled crosslinking in resilin makes it reasonable to suggest, that chitin-protein cross-linking, probably by histidine and aspartic acid (Hackman, 1960), could also occur in the deposition zone. Unless the chitin is actually assembled on the protein chains, as suggested by Fraenkel and Rudall (1947), it is assumed that any orientation must take place before the cross-linking of the protein to the chitin. Under these conditions there could then be an opportunity for mutual orientation effects between the

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chitin and protein molecules, and between the protein molecules themselves, which would then be stabilized by cross-linking and hydrogen bonding. Better mutual orientation of chitin and protein would lead to a decrease in separation and an increase in number of prospective reaction sites. It may even be that extensive cross-linking cannot occur until the chitin and protein chains are optimally orientated. The ultimate forces involved in determining the relative orientations of large molecules in aqueous environments are London-van der Waals forces and coulombic electrostatic charges. According to the DebyeHuckel theory ( I 923) net electrostatic repulsion or attraction varies with the ionic concentration of the medium (Fig. 38). Raising the ionic concentration lowers the electrostatic repulsion between like charged groups, and also the attraction between unlike charged groups. The final mutual orientation of chitin and protein would depend upon a very complex steric interaction of the above kind of forces, and would be dependent upon the ionic concentration of the medium in the deposition zone prior to cross-linking. Since dissociation of polar

1.5 x

lo-"

u)

-P

C

0

-

I.Ox

10-12

0 3

6

c

V

E

P

+ W

c

c .-

z

3.5 x lo+

.z 2.

P 0 C

-

2

0 .c

al c c

c

a"

0 c

" e

-0.5 x

c

0

K = 10'

c

W

z K =

3 x lo6/

4

Separation distonce

S

FIG.38. Hamaker curves showing the influence of electrolyte concentration, K , on the total potential energy of interaction (London attraction and static electric repulsion, the latter being modified by Debye-Huckel double-layer screening) (after Kruyt, 1952).

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groups is dependent upon pH, the electrostatic forces would also vary with changes in hydrogen ion concentration. We may note here the possible significance of histidine in cuticular proteins. Histidine is the only amino acid, with the exception of the N-terminal amino groups, with a pK value in the region of physiological pH, and could thus contribute to a charge on the protein. On contact with chitin, the extra amino group could form an acid amide with the acetyl group of the chitin side chains as proposed by Hackman (1959). We must now attempt to account for the charges on the protein and chitin molecules. In the case of the arthropodin complex, glutamic and aspartic acids and the basic amino acids (histidine, lysine and arginine) are all present to varying extents in various species (Hackman, 1960). In the cockroach, Periplaneta americana, the relative concentrations of amino acids in the cuticle appear to vary throughout the intermoult period (Lipke, et al. 1965); the significance of this is unknown. Poly-N-acetylglucosamine sensu strict0 is not a charged polysaccharide. Thus the charges on native chitin depend upon the possibility that not all of the glucosamine residues are acetylated, since each non-acetylated glucosamine would carry a positive charge. There is now a considerable amount of evidence in support of this. Giles et al. (1958) performed elemental analyses on lobster chitin which had been very gently deproteinized so as to avoid artificial deacetylation. They then compared the quantity of C, H and N with the theoretical quantity in poly-N-acetylglucosamine, and concluded that the best estimated composition was : N-Acetylglucosamine 82.5% (by weight) Glucosamine 12.5% Water 5.0'4 Waterhouse et al. (1961) extracted glucosamine and N-acetylglucosamine from crab cuticle with a chitinase preparation which showed no deacetylase activity, and found up to 10% glucosamine in native chitin. These facts suggest that every sixth glucosamine residue along the chitin chains may be non-acetylated. Rudall (1963) suggests that this may explain the 31 A repeat along the polymer chains which was revealed by his X-ray diffraction analysis of Hymenopteran and Orthopteran cuticle and Aphrodite chaetae (31 A = 6 x b axis repeat period = 6 x 5.14 A = 30.84 A) (Fig. 1).*

* Hackman and Goldberg (1965) found one glucosainine for every 4 acetylglucosainine residues in fl-chitin from cuttlefish, and 2.2 glucosatnines for e v e y 3.2 acetylglucosamines in Loligo chitin, based upon results from deuteration studies, enzymic digestion and infrared absorption spectra.

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There is further evidence for the presence of glucosamine in native chitin from the PAS test, which reacts with terminal groups of chitin chains (Jeanloz and Forchielli, 1950) and with non-acetylated residues (Brunet, 1952). Endocuticle is weakly PAS positive presumably varying with the number of short chain molecules and with the degree of non-acetylation. The zone of deposition is strongly PAS positive (Taylor and Richards, I965), again perhaps indicating short chain lengths and easily accessible non-acetylated glucosamine residues. Taylor and Richards interpret this as indicating that the zone of deposition or subcuticle is (1) the region into which cuticle precursors are secreted, (2) the region where polymerization of N-acetylglucosamine residues occurs and (3) the region where chitin-protein crosslinkages are formed. All of this is compatible with the above orientation hypothesis. Marchessault et al. ( 1 959) showed that alkaline purification of chitin led to a change in pH to 3.5, as more and more unacetylated groups were created or uncovered, and their amine groups complexed with protons to form NH,+. It is suggested then that the degree of unacetylated glucosamine may give a positive charge to chitin chains and that this is instrumental in determining their orientation when the ionic composition of the deposition zone is varied by the cells. (In this connection it is interesting to note that cellulose in plants contains a small number of uronic acid side chains.) Orientation control could thus be effected by : 1 . p H changes in the extracellular environnient 2. Ionic changes in the extracellular environment

A rhythmical ion pump could, for instance, control lamellogenesis frequency. Divalent ions would have a greater screening effect on polar groups than monovalent ions. This might, in fact, provide an explanation for the lack of orientation of protein chains in the chitin-free oothecae of cockroaches (Pryor, 1940), for large quantities of C a + + ions are known to be present in these structures. Hackman and Goldberg (1960), for example, found 6.5% calcium in the ootheca of Periplaneta umericana. Scudamore (1 947) has shown the epidermis which secretes crayfish gastroliths is capable of C a + ion deposition on a daily basis, incorporation occurring nocturnally. If such a mechanism operated in the epidermis which secretes insect cuticle it could provide an explanation for the daily orientation changes. The biological significance of the inorganic ions present in insect cuticles is so far unknown (Hackman, 1964). Travis (1963) suggests that the continued increase in ash and +

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mineral content, which occurs in parallel with the mineralization of crustacean cuticle and gastroliths, is evidence that the epidermal cells control the calcium, carbonate, phosphate and local pH changes in the extracellular environment. Glimcher (1960) has made similar suggestions concerning the environment surrounding vertebrate collagen fibrils and their combined effect upon induction of mineralization 3. Rhythmical water absorption. This could also control lamellogenesis frequency according to the following scheme (Fig. 39).

/

Secretion phase : polymerization by condensation

Extracellular water concentration falls : ionic concentration rises, causing orientation change

\

Extracellular water concentration rises : ionic concentration falls, causing orientation change

phase FIG. 39. A possible cycle for chitin lamellogenesis control.

4.A combination of the above factors Evidence from several sources suggests that epidermal cells can control the state of hydration of cuticle. Experiments with cockroaches (Beament, 1964) show that the rate of water uptake through the hard regions of the cuticle is higher than through the softer regions, probably varying inversely with their respective states of hydration. That these regional differences are actively maintained by the cells was shown by the fact that in anaesthetized cockroaches the rate of water uptake was constant all over the skeleton. Winston and Nelson (1965) have recently demonstrated active integumental control of water loss in the clover mite Bryobia pruetiosa (Acarina, Tetranychidae). Bennet-Clark (4 96 1) measured the degree of extension of samples of Rhodnius cuticle by various loads

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after different times and at controlled pH values. He suggests that hydration and extensibility follow dissociation of the COOH groups of the protein at low pH values, and that the control of cuticular plasticity in the bug following a blood meal is brought about by a change in the hydration of the cuticle. It is worth noting here that nonlamellate locust endocuticle produced in constant day conditions is noticeably more dry and brittle than control reared cuticle (A. C. Neville, unpublished). These results indicate that the epidermal cells can exert regional control over the hydration of the cuticle, and hence also of the deposition zone. Such a mechanism, if coupled with a temporal control system, could influence both the primary orientation and also the secondary reorientation of chitin by changing the ionic environment of the cuticle. The defective crystallinity of cellulose microfibrils in plants is due largely to the occlusion of water molecules liberated during crystallization (Frey-Wyssling, 1957). In view of the evidence, discussed in Section V A above, that chitin polymerization is extracellular, and of the suggested involvement of hydration control, water removal may play an important part in cuticle morphogenesis (B. L. Gupta, personal communication). It is therefore not surprising to find a well developed brush border of micro-villi in epidermal cells secreting insect cuticle (Locke, 1961, 1964). Such brush borders are typical of water and ion absorbing cells, e.g. vertebrate renal tubule cells; absorptive region of malpighian tubules in insects (Smith and Littau, 1960); absorptive epidermis of ventral gland in Podura aquatica (Noble-Nesbitt, 1963a,b). Control of ionic concentration across the deposition zone would also overcome the difficulty concerning the microvillate epidermal surface, which as Locke (1964) notes, could not act as a specific template for chitin architecture. The well known in virro reconstitution of SLS and FLS collagen from tropocollagen (Randall et al., 1955; Schmidt, 1956) can be controlled by regulation of pH and ionic concentration. The resulting periodic banding patterns represent sophisticated examples of macromolecular orientation. There is some evidence for the operation of long-range forces in chitin and cellulose orientation in vitro. Marchessault et al. (1959) prepared some liquid crystals of chitin and cellulose which showed local parallel orientation of crystallites. Low angle X-ray measurements showed that the interparticle distance was about 400 8, for a 15% gel, varying as the square root of the concentration. They postulated long range forces or entropy effects to explain the parallel alignment.

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The recent work by Ben-Hayyim and Ohad (1965) on synthesis and orientation of cellulose by the bacterium Acetobacter xylinum, demonstrates the existence of an electrostatic orientation mechanism of the kind proposed above for insect chitin. This bacterium can be made to incorporate the cellulose which it produces extracellularly, normally in a randomly oriented manner, into a medium containing other polysaccharides by a process of co-crystallization. The resulting fibrils are oriented in parallel, the direction changing from layer to layer, if the cellulose is incorporated into a charged polysaccharide (carboxymethylcellulose (CMC) or phosphomannan); but if, however, they are incorporated into a non-charged one (levan) then they are randomly oriented. The degree of orientation with CMC was reduced by lowering the pH, which acted by reducing the charge on the carboxyl groups of the CMC. Similarly, aggregation of fibrils was delayed by electrostatic repulsion between the charged carboxyl groups when CMC was employed. This aggregation time was experimentally decreased by reducing pH or by increasing the ionic strength of the medium. It is of some interest to note, therefore, that in a polysaccharide system, closely similar to that of chitin, orientation of fibrils into parallel layers (which change direction from layer to layer) can be brought about by regulating the ionic composition of the medium into which a number of unco-ordinated bacterial cells are cumulatively secreting. Hayyim and Ohad suggest that charge interaction could be part of the mechanism of orientation of cellulose fibrils in plant cell walls, and that specific patterns could be due t o the presence of different polysaccharides.

VI. CONCLUSION The above account has shown how changes in periodic orientation during cuticle morphogenesis may be structurally advantageous. It has also been suggested that the cellular control of those changes is effected by regulation of physical factors in the extracellular environment in which deposition, charge interaction between chitin and protein, orientation and chitin-protein cross-linking occur. It is further suggested that the temporal control (daily or microcycle) which regulates the ordering of the orientation changes, controls the structural sequence in the fabrics produced. Because of their easily accessible single-layered epidermis, insects should provide suitable material for testing the hypothesis of ionic and electrostatic control of orientation in uiuo. It is to be hoped that such research may help to increase our knowledge of some of the general principles underlying”macromolecular morphogenesis. ~ ~ + A . I . P4.

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Taylor, R. L. and Richards, A. G. (1965). Integumentary changes during moulting of arthropods with special reference to the subcuticle and ecdysial membrane. J. Morph. 116, 1-22. Thor, C. J. B. and Henderson, W. F. (1940). The preparation of alkali chitin. Am. Dyestuff Reptr. 29, 461-464. Timoshenko, S. (1951). “Theory of Elasticity”. McGraw-Hill, New York. Travis, D. F. (1963). Structural features of mineralization from tissue to macromolecular levels of organization in the Decapod Crustacea. Ann. N . Y. Acad. Sci. 109, 177-245. Travis, D. F. and Friberg, U. (1963). The deposition of skeletal structures in the Crustacea. VI. Microradiographic studies of the exoskeleton of the crayfish, Orconectes virilis Hagen. J. ultrastruct. Res. 9, 285-301. Verhoeff, K. W. (1926-32). Diplopoda. Bronns KI. Ord. Tierreichs 5 (2), 3. Vignon, P. (1901). Recherches sur les epitheliums. Archs. Zool. exp. gdn. (sCr. 3) 9,371-715. Waterhouse, D. F., Hackman, R. H. and McKellar, J. W. (1961). An investigation of chitinase activity in cockroach and termite extracts. J. Insect Physiol. 6, 96-1 12. Weis-Fogh, T. (1959). Elasticity in arthropod locomotion : a neglected subject, illustrated by the wing system of insects. Proc. XVth. int. Congr. Zool., 393. Weis-Fogh, T. (1960). A rubber-like protein in insect cuticle. J. exp. Biol. 37, 889-907. Weis-Fogh, T. (1963). Resilin, a rubber-like protein, and its significance. In “Aspects of Protein Structure” (G. N. Ramachandran, ed.), pp. 337-341. Academic Press, New York. Wigglesworth, V. B. (1930). The formation of the peritrophic membrane in insects, with special reference to the larvae of mosquitoes. Q. JI microsc. Sci. 73,593-616. Wigglesworth, V. B. (1956). Formation and involution of striated muscle fibres during the growth and moulting cycles of Rhodnius prolixus (Hemiptera). Q. JI microsc. Sci. 97,465480. Winston, P. W. and Nelson, V. E. (1965). Regulation of transpiration in the clover mite Bryobia praetiosa Koch (Acarina: Tetranychidae). J. exp. Biol. 43, 257-269. +

Addenda Page 214: Most crystalline polymers crystallizefrom solution in the form of 100 A thick

lamellae, in which the molecules are folded back and forth on themselves (Geil, 1963). A folded-chainsingle molecule model for cellulose microfibrils has been proposed by Manley (1964). Page 225: The formation of parabolic moirC patterns from laminate spherulites in which each consecutive lamina of parallel oriented molecules progressively changes orientation, is known from polymer systems (reviewed by Geil, 1963). REFERENCES

Geil, P. H. (1963). Polymer single crystals. “ Polymer Reviews ”,Vol. 5. New York. Manley, R. St. John (1964). Fine structure of native cellulose microfibrils. Nature, Lo&. 204,1155-1157.

The Biochemistry of Sugars and Polysaccharides in Insects G . R. WYATT Department of Biology, Yale University, New Haven, Connecticut, U.S.A. I. Introduction . 11. The Occurrence of Sugars in Insects . A. Glucose and reducing substances . B. Trehalose C. Sugar content of insect hemolymph . D. Sugar content of whole insects and insect tissues . 111. Intestinal Absorption and the Physiology of Hemolymph Sugar Levels . A. Absorption from the gut . B. Regulation of blood sugar . IV. Biosynthesis and Utilization of Sugars . A. Glucose . B. Use of monosaccharides other than glucose . C. Biosynthesis of trehalose . D. Cleavage and use of trehalose . E. Physiological roles of trehalose and trehalase . F. Dormancy and the properties of trehalose . V. Glycogen . A. Glycogen in insects . B. Accumulation and conversion during growth and metamor. phosis C. Glycogen in insect flight . D. Metabolism of glycogen . VI. Hormonal Effects on Carbohydrate Metabolism VII. Glycoproteins and Chitin . A. Glycoproteins in insects . B. Metabolism of chitin VIII. Glycerol and Sorbitol . References .

287 289 289 289 29 1 295 297 297 299 30 1 301 302 304 309 317. 324 325 325 327 329 329 336 340 340 341 345 347

I . INTRODUCTION

In their carbohydrate metabolism, insects make use of substances, enzymes and operational principles that are known from other living groups. Yet the ways in which these fit together, particularly with respect to tissue interactions and regulatory mechanisms, forms a distinctive system of which the main features are just now being 11.

287

.

288

G . R. W YAT T

discerned,and which differsin severalimportantrespects from the pattern well known in mammals and other vertebrates. Our knowledge of carbohydrate metabolism in other invertebrate groups is too fragmentary for one to judge to what extent the insect pattern is unique, but it is likely that its features are shared in varying degree. Occupying a central place among these is the presence of trehalose as the predominant blood sugar. Its distribution and the relations of its metabolism to the mechanisms of nutrient absorption, storage and supply to the tissues, will be discussed in some detail and will occupy a major part of this review. Characteristic of insects and other arthropods is the presence of chitin as a structural component of the exoskeleton, which in its deposition and resorption interacts closely with the metabolism of other carbohydrates. The mucopolysaccharidesof insect hemolymph and tissues have only recently begun to be investigated. A metabolic process apparently peculiar to insects and some other arthropods is the bulk conversion of glycogen to glycerol and sorbitol. Glycogen, chief reserve carbohydrate in insects as in other animals, appears to be metabolized by familiar pathways, but under the guidance of a special insectan group of hormones. These subjects will be treated, but without claim to complete coverage of the literature. In these and other aspects of insect biochemistry, of course, the special demands of the cycle of moulting and metamorphosis loom large. It is hoped that this rather specialized review will complement others which have appeared on carbohydrates and their metabolism in insects in recent years. Chefurka (1964, 1965) has treated the subject broadly, while Kilby (1965) has discussed carbohydrate metabolism in insect fat body and Sacktor (1964) has covered aspects related to muscle contraction. Conparative aspects of carbohydrate digestion are treated by Barrington (1962) and, more recently, House (1964a, b) has reviewed both digestion and nutrition in insects. Discussion of chitin and its metabolism may be found in a monograph by Jeuniaux (1963), and there exist several reviews on the chemistry of insect cuticle (Richards, 1958; Rudall, 1963; Hackman, 1964). The metabolism of trehalose in animals, including insects, has recently been reviewed in Russian by Lukomskaia (1964). It is fortunate for the study of the biochemistry of relatively small animals, such as insects, that technical advances now make possible precise experimentation with very small amounts of tissue. This is especially important since the biologically sigdcant question of the interactions of functionally discrete tissues has been little investigated in insects until very recently. The use of whole-animal homogenates has

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often obscured tissue differences. Such preparations must sometimes be used as a ready source of suflicient material, as for enzyme purification. But it is important to supplement such work with experiments to assign the tissue source of what is found. Although insects have provided the material for several well-known original biochemical discoveries, the investigation of the biochemistry of this animal group has in general lagged far behind that of micro-organisms on the one hand and vertebrate animals on the other. Recently, an infusion of experimental methodology and critical standards from the better-developed regions of biochemistry has been taking place, and this process should assist our taking advantage of the biological peculiarities of insect material. 11. THEOCCURRENCE OF SUGARS I N

INSECTS

A. GLUCOSE A N D R E D U C I N G SUBSTANCES

In insects, as in other animals, glucose has a central place in carbohydrate metabolism. In the majority of species that have been examined, however, the amount of free glucose is quite small. As has been repeatedly noted (e.g. Vaney and Maignon, 1906; Hemmingsen, 1924; Howden and Kilby, 1960), the substances in insect blood which react in the traditional clinical sugar assays that measure “reducing value ” are chiefly not sugars. Several amino acids, notably tryosine and proline, appear to be responsible for much of the non-sugar reducing substances, which are frequently at high levels (Heller and Mochnacka, 1951; Howden and Kilby, 1961). An estimate of fermentable sugars can be obtained by measuring reducing value before and after a brief fermentation by washed yeast, but, where the difference measured is a small fraction of the total the error is likely to be considerable. True carbohydrate levels are now best measured by more specificchemical methods (such as the anthrone reaction for total glucose derivatives), specific enzymic assays (such as glucose oxidase supplemented with trehalase) and several types of chromatography. B. TREHALOSE

The most characteristic sugar of insect hemolymph is trehalose, a symmetrical disaccharide of glucose (a-D-glucopyranosyl-a-D-glucopyranoside) which is non-reducing by virtue of tlie anomeric carbon atoms of both glucose moieties being bound in the glycosidic linkage:

290

G . R. WYATT

OH Much of the earlier work on trehalose and its metabolism, especially in yeast, is reviewed by Myrback (1949), and its chemistry has recently been treated by Birch (1963). In the Paris Universal Exposition of 1855 there was displayed among a collection of oriental pharmaceutical products a sample of trehalu which apparently originated in Syria, where this material had long been an article of commerce (Guibourt, 1858; Hanbury, 1859). The chemist Berthelot (1858a, b, 1859) obtained some of this material and showed that, along with polysaccharides, it contained a high proportion of a sugar which he named trehalose. Berthelot commented upon the close similarity of his trehalose to mycose, a fungal sugar obtained from rye ergot (Wiggers, 1832; Mitscherlich, 1858); the two were subsequently shown to be identical, but the more colourful name stuck. The contemporary descriptions of trehala as the shells or cocoons (coques) of a weevil (Larinus nidificans)found in masses on the stems of its host plant, the thistle-like Echinops persicus, left no doubt that it was an insect product. This is supported by the fact that the sugar trehalose is not known to be produced by flowering plants. Trehala is later discussed and depicted by Pierce (1915), and trehalose has been isolated from another “desert manna” consisting of dried excretions of scale insects by Leibowitz (1943, 1944). The association of trehalose with an insect, however, was generally forgotten and it came to be regarded as a sugar characteristic of lower plants (Myrback, 1949). Just one hundred years after the exhibition of trehalu in Paris, after a number of indications that insects contained some unrecognized carbohydrate (see Wyatt, 1961), trehalose was re-discovered in this animal group as a result of almost simultaneous work in three laboratories (Wyatt and Kalf, 1956, 1957; Howden and Kilby, 1956; Evans and Dethier, 1957). It was found that insect hemolymph often contains so much trehalose that the sugar can easily be obtained from it in crystalline form. Despite isolated scepticism (Hansen, 1964), crystalline trehalose and derivatives have been prepared from insect blood by yet

B I O C H E M I S T R Y OF S U G A R S A N D P O L Y S A C C H A R I D E S

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other workers (Randall and Derr, 1965), and this sugar has been identified in every insect species that has been appropriately examined, although it could not be detected in some developmental stages of some species. At the same time as its re-discovery in insects, trehalose was discovered independently in Ascaris (Fairbairn and Passey, 1957), and this was followed by its identification in many other invertebrate animals belonging to several phyla (Fairbairn, 1958). Trehalose occurs generally in fungi, in many algae (see Myrback, 1949) and in occasional pteridophytes (Selaginella and Botrychium ;Kandler and Senser, 1965). It has never been identified, however, as a product of the highest groups of plants or animals, the flowering plants and the vertebrates.* C . S U G A R C O N T E N T OF I N S E C T HEMOLYMPH

In Table I are compiled data from the literature on sugar levels in insect hemolymph. Some of these values have appeared in previous compilations (Altman and Dittmer, 1961; Wyatt, 1961; Florkin and Jeuniaux, 1964). In accordance with traditional usage, results are expressed in units of weight, but it is encouraging to find the biochemically more meaningful molar units appearing in some recent papers. Sugar levels are generally high compared with those found in vertebrates-it is usual to find more than 0.5% sugar in insect blood and there are several records of more than 5%, the highest apparently being that recorded by Ehrhardt (1962) for the aphid Megoura viciae (up to 8.1% total sugar in the hemolymph). The most usual picture is a relatively low level of glucose along with a higher one of trehalose, but there are many exceptions. Thus, in larval hemolymph of the blowfly, Phormia regina (Evans and Dethier, 1957), trehalose could not be detected, although it appears at pupation and is abundant in the adult. Similarly, in the sphingid, Celerio euphurbiae, trehalose first appears in the hemolymph at the cessation of larval feeding (Mochnacka and Petryszyn, 1959). In other cases, atypical carbohydrate content in hemolymph appears to reflect specialized diet. Thus, the abundant fructose in larval hemolymph of the horse bot fly, Gastrophilus intestinalis (Levenbook, 1950), is probably attributable to the high levan content of the host’s food. Small amounts of fructose and sucrose found in blood of the silkworm (Wyatt et al., 1956) and the aphid, Meguura viciae (Ehrhardt, 1962), probably derive from plant sap. The influence of diet on hemolymph

* In an isolated report of trehalose from the sedge, Carex (cited by Myrback, 1949), the sugar was found in exudates apparently produced by some pathogenic organism and not in the sedge itself (von Lippmann, 1912).

TABLE I

h,

W

h,

Carbohydrate levels in insect hernolymph* ~~

Content in hemolymph (mg/ml)t Order and species

Orthoptera Gryllus domesticus

Stage

A

Leucophaea maderae Locusta migratoria

A

L. migratoria

A

Glucose

Trehalose

0.8

3-5

0.6

5.8-7.8

Glycogen ''$

0.25 0-20

14-5f 3.5

4*0(1*8-7.2)

present

14.0(9*9-18*1) 8.8 k 0.8

Schistocerca gregaria L, V

0-2

2-20

A, P

0.24(0.06-0.40)

7.0(5*5-8.5)

L A,

13-20 10-11

21-36 58-71

P

Other sugars (mglml); remarks

Reference

G estd. by difference; Nowosielski and T shows circadian Patton, 1964 rhythm G estd. as fermentable Todd, 1958 reducing power Humphrey and Fructose 0.35 Robertson, 1949 Total sugars about 20; Hansen, 1964 maltose, cellobiose and sometimes fruo tose present; vary with diet Randall and Derr.

2.8(2.141)

Melanoplus A, P differentialis Periplaneta americana A P. americana A

S. gregaria Homoptera Megoura viciae

"

1965

0.3-05

4-8

Treherne, 1960 T falls during &gbt Polacek and Kubista, 1960 Levels rise during in- Howden and Kilby, star, vary with diet. 1956, 1960 Treherne, 1958a. b G estd. by difference; Ehrhardt, 1962 traces of fructose and sucrose present

Lepidoptera Antheraea pernyi

L,V

Antheraea polyphemus L and P Bombyx mori L,v B. mori

L, IV, v

B. mori B. mori

L, P

Celerio euphorbiae

L

C . euphorbiae

P

Deilephila elyenor

P

Galleria mellonella

L

Hyalophora cecropia

v

T rises during instar

3.2-8.8 0.01-0.03

10-14 3-1-5.2

0.14

1-3

0.02-0.05

4.56.0 2.0

0.4

T absent in feeding larva

0-16 0.7-20

5-13

I

9-19

0408

0.2

15

0.4

L, V

0

12-19

Leucania separata

P P L, V, VI

0.02-0.1 0-0.1 0-2-0-9

3-7 10-15 1-5

Samia Cynthia

P

0 0-0.03

2-5 5-1 5

S. c. ricini

L,v

Sphinx l i g h r i

P

24

G rises to 0.4 at moult; fructose 0.015; G-6-P 0.4-1.2 T rises and falls at moults T falls during starvation

0-2-0.5

0

G estd. as fermentable reducing power; T falls t o 1.3 in developing adult T 2-12 in developing adult

Smolin, 1960; Egorova, 1963 Wyatt andKalf, 1957 Wyatt et al., 1956; 0 Wyatt and Kalf, n 1957 Duchateau-Bosson et al., 1963 Saito, 1963 Wyatt et al., 1956; Wyatt and Kalf, 1957 Mochnacka and Petryszyn, 1959 Heller and Mochnacka, 1951 ; Mochnacka and Petryszyn, 1959 Duchateau and Florkin, 1959 Wyatt and Kalf, 1957 Wyatt and Kalf, 1957; Wyatt, unpublished 9

Maximal just before spinning Pupae in diapause Developing to adults G and T maximal just Liu and Feng, 1965 before pupation Pupae in diapause Wyatt, unpublished T maximal just before Chang et al., 1964; spinning Liu and Feng, 1965 Duchateau and Florkin, 1959

z

n n X

s: w

TABLE I-(contd.) N

Content in hemolymph (mg/ml)t Order and species

Stage G 1u cose

Trehalose

“

Glycogen ”$

Coleoptera Chalcophora mariana L

52

0

Dytiscus marginalis

A

5-7

0-0.1

Ergates faber

L

33

0

Hydrous piceus

A

0.05-0.3

3-5

0

Tenebrio molitor Hymenoptera Anihophora sp.

L

1.2

Apis mellifera

A

Dbrion hercyniae Diptera Gasirophilus intestinalis C . intestinalis Phormia regina P . regina

L

Other sugars (mglml); remarks

Duchateau and Florkin, 1959 Duchateau and Florkin, 1959 DuChdteaU and Florkin, 1959 G estd. as fermentable Florkin, 1937; reducing power Duchateau and Florkin, 1959 Marcuzzi, 1956

65 11-14

L

0.3

L, 111

0-1

Fructose 8-10

6-12 8

0.3 0.1-0.2

P L

0.7-1.2

0

0.3

A

0.3-6

1.2-30

0.7

Reference

Duchateau and Florkin, 1959 Czarnovsky, 1954; Duchateau and Florkin, 1959 Wyatt and Kalf, 1957

Fructose 2G-2.8

Levenbook, 1950

Fructose 0.2-1.0

Levenbook, 1947 Evans and Dethier, 1957 Evans and Dethier, 1957

T falls during flight

* Analyses by chromatographic, enzymic and specific chemical methods are listed; most of those based on reducing power are omitted. L, larva (roman numeral designates the instar); P, pupa; A, adult; G, glucose; T, trehalose. 7 For trehalose (mol. wt. 342) I mglml = 2.92 mM; for glucose (mol. wt. 180) 1 mg/ml = 5.56 mM. $ “Glycogen” is not identified as such, and usually represents material resistant to alkali and insoluble in alcohol, or immobile on paper chromatograms, determined with anthrone reagent.

n P

< ~

ci 4

BIOCHEMISTRY OF SUGARS A N D POLYSACCHARIDES

295

carbohydrate composition has been shown experimentally. Hansen (1964, in a paper which also reports the isolation of crystalline maltose and cellobiose from locust hemolymph) presents evidence that fructose is present in the blood of locusts fed on fructose-rich food (pears) but not in locusts fed on wheat or Brussels sprouts. The honey bee (Apis rnellgera) is exceptional in the high content of free reducing sugars as well as trehalose in its blood; thus, adult bee hemolymph contains glucose, fructose and trehalose each at a level of the order of magnitude of 1% (Beutler, 1936; von Czarnovsky, 1954; Duchateau and Florkin, 1959). Recently, Maurizio (1965) has reported extensive analyses of sugars in bee blood by paper chromatography, which was applied to numerous samples from bees of different castes in different seasons and on different diets, natural and artificial. Quantitative results are given for each sugar as its percentage of the total sugar revealed on the chromatograms by anisidine hydrochloride reagent. Trehalose was recognized but not estimated. It is a pity that the results of this meticulous work cannot be expressed in standard units of concentration. In addition to glucose (0.3-56% of total sugars detected) and fructose (28-99%), which were universally present, certain samples contained smaller but significant amounts of sucrose (0-23%), maltose (0-19%), and the trisaccharide fructomaltose (L-maltosyl-fla-fructofuranoside, also known from honeydews; Gray and Fraenkel, 1954; 0-873. The relative proportions varied greatly in different groups of bees. Some similar results, with more fructose than glucose, were reported for a bumble bee (Bornbtls).Interesting results, which show the non-specificity of sugar absorption, were obtained by feeding caged bees on pure sugar solutions; after 24 h on fructose the blood reducing sugar was 98-100% fructose; after 24 h on glucose, maltose or trehalose, the blood revealed only glucose (with the exception of one group, in which some maltose persisted as such; note also that the analytical method was not sensitive to trehalose). After 6 h on galactose (which was toxic), only galactose was found in the blood. After feeding the trisaccharide melezitose, (a-D-glucopyranosyl-(1+-3)-fl-~-fructofuranosyl-(2+1)-a-D-glucopyranoside), glucose, fructose and melezitose itself appeared in the blood. D. S U G A R C O N T E N T O F W H O L E I N S E C T S A N D I N S E C T TISSUES

Some analyses have been performed on the sugar content of whole insects. In Antheraea pernyi, trehalose is present throughout the life

296

G. R. W YAT T

cycle, and undergoes a steady increase in level from the egg (0.026% of dry weight) to the pupa (up to 2.6%), then a decline in development to the adult (Egorova and Smolin, 1962a). The presence of trehalose within eggs, and its quantitative changes during embryonic development, have been demonstrated in Bombyx mori (Dutrieu, 1961a, b; Yamashita, 1965) and Melanoplus diflerentialis (Randall and Derr, 1965). In certain overwintering insects, the total sugar levels are extraordinarily high. Prepupal larvae of a sawfly, Trichiocumpus populi, which, after pre-freezing at -20 to -30" can survive 2 h in liquid nitrogen, contain sugar equal to 5-9% of their fresh weight, and this is stated to be 97% trehalose (Ashahina and Tanno, 1964). Certain adult solitary bees (Cerutinu frauipes and C . japonicu), which also overwinter, contain still greater sugar levels: l0-15% of body weight, comprising 45-77 mg/g fructose, 3048 mg/g glucose and only about 4 mg/g trehalose (Tanno, 1964) (the trehalose level is thus similar to that of many non-overwintering insects). These bees cannot withstand freezing, but there is an indication that the high sugar content lowers the supercooling point and confers frost-resistance in this way. They are stated to contain neither polyols (see Section VIII) nor glycogen. The tsetse fly (Glossinu spp.) has been examined because of the possible importance of its sugars in parasitism by trypanosomes, and the levels found by enzymic assay were exceptionally low: in whole flies and in salivary glands less than 0.04 mg/g trehalose and less than 0.06 mg/g glucose, while intestinal tissue contained up to 0.2 mg/g trehalose (Geigy et ul., 1959). Further examination by chromatography of alcoholic extracts of adult tsetse flies revealed arabinose and traces of sucrose (Wyss-Huber et al., 1961). This appears to be the only report of free arabinose from an insect, though it does occur in insect glycoproteins (see Section VII). Information on sugar levels in defined insect tissues is scanty, which is regrettable in view of the importance of such data for the interpretation of intermediary metabolism. Its paucity is due in part to experimental difficulties in assuring complete rinsing of tissue while avoiding leaching of cell contents, and in avoiding interference in the analysis by tissue components. It appears that the internal sugar levels tend to be much lower than those in the surrounding hemolymph. Mochnacka and Petryszyn (1959) examined separately the hemolymph and the combined bled tissues of Celerio euphorbiue, a sphingid in which trehalose is apparently restricted to the pupal stage. In the early pupa, when blood trehalose was in the neighborhood of 10 mg/ml, none

BIOCHEMISTRY OF SUGARS AND POLYSACCHARIDES

297

could be detected in the tissues. Later, during diapause, trehalose appeared in the tissues at 2-3 mg/g (being still about 10 mg/ml in the blood), then during adult development trehalose declined and disappeared, first from the tissues and then from the hemolymph. In Hydophora cecropia, trehalose has been estimated in tissue water of larval fat body (a site of active trehalose synthesis) as about 15 m~ (G. R. Wyatt, unpublished), while the level in hemolymph was about 50 mM. Other determinations of trehalose, glucose and various intermediates of carbohydrate metabolism have been performed in relation to the analysis of metabolic regulation in flight muscle (which will be further discussed in Section IV). In flight muscle of Locusta migratoria, trehalose has been reported as 4-21 pmolelg (1-4-7.2 mg/g) and glucose as 0.8-5.6 pmole/g (0.14-1-0 mg/g) (Bucher and Klingenberg, 1958). In the blowfly, Phormia regina, trehalose was found in adult fight muscle at about 0-3 mg/g, compared with 10-30 mg/ml trehalose and 2-3 mg/ml glucose in the hemolymph (Clegg and Evans, 1961). The levels in hemolymph being known, relatively low tissue sugar levels can also be inferred from the recent data of Sacktor and Wormser-Shavit (1966) on whole thoraces of P . regina: trehalose, 2-7 prnolelg (0.7-2-4 mg/g) ;glucose, 3.6-7-6 pmole/g (0-6-1-4mg/g).

111.

ABSORPTION A N D THE PHYSIOLOGY HEMOLYMPH SUGARLEVELS

INTESTINAL

OF

A . A B S O R P T I O N FROM T H E G U T

In 1906, Vaney and Maignon concluded from experiments that in the silkworm " le sucre est detruit, au cours de la digestion, au niveau de Epithelium intestinal, ou immidiatement B son arrivee dans le sang." As to the mechanism of absorption, however, nothing was known until quite recently (see Waterhouse and Day, 1953). The problem has been studied by Treherne (1957) in a preliminary fashion, in the cockroach, PeripZaneta americana, and in detail in the locust, Schistocerca gregaria (Treherne, 1958a-c). The method was to !ill the gut in viuo or in vitro with a solution of [14C]sugarand the nonabsorbed dye Amaranth, then to sample portions of the gut contents and measure the ratio of radioactivity to colour. In the locust, absorption occurred chiefly from the mid-gut caeca and to a lesser extent from the ventriculus. The rate of loss of [14C]glucose,when introduced into the gut at different low concentrations, was a constant proportion of the amount present, and was anaffected by poisoning with cyanide

298

'

G . R. W Y A T T

and iodoacetate. In vivo, glucose did not accumulate in the hemolymph, but was rapidly converted to trehalose, except when the amount administered was so high as to saturate the mechanism for trehalose synthesis temporarily. Mannose and fructose were absorbed at lower rates, and appeared in the hemolymph, being more slowly converted to trehalose. These observations are interpreted as showing that sugar absorption in the locust is accomplished by passive diffusion, facilitated by conversion of the absorbed sugar into trehalose. Injected glucose was converted to trehalose at the same rate as that from the gut; thus, this synthesiswas independent of absorption.The paradox of rapid absorption of radioactive glucose from the gut when its concentration (2 mM) was scarcely higher than that in the hemolymph (1.3 mM) was resolved by experiments involving injection of [14C]glucose into the hemocoele. These indicated that exchange of glucose took place across the gut wall, and with low glucose levels in the gut there was probably no net absorption. Recent experiments with insect intestines in vitro have also given results supporting passive diffusion as the mechanism of sugar transport. Randall and Derr (1965) found that everted intestines of the grasshopper, Melanoplus diflerentialis, suspended in glucose-containing medium failed to concentrate the sugar internally, contrary to the results of others who had applied the same technique to mammalian intestinal segments. Shyamala and Bhat (1965) perfused mid-guts of the silkworm, Bombyx mori, in an ingenious apparatus and found with the use of [14C]glucosethat the sugar passed in both directions (0.346 ~ in a manner indicative of diffusion. pmole/h at 2 m concentration) Its passage was unaffected by the presence of dinitrophenol, and there was no evidence of active transport. In the same experiments, water was transported toward the lumen in a process that was inhibited by dinitrophenol and therefore dependent on metabolic energy. In similar perfusion experiments with the mid-gut of adult Phormia, Gelperin (1966) found no evidence for active transport of sugars. Fructose passed through the gut wall at the same rate as glucose, and the movement of both was unaltered by several metabolic poisons. The capability of the insect gut for active transport of water and of inorganic ions is well established (Treherne, 1965), but in contrast to vertebrate animals insects seem not to use such transport for sugar absorption. The occurrence of sugars in insect hemolymph is also consistent with relatively non-specific absorption by diffusion. Thus, unusual sugars which are present in the digestive tract may appear in the blood, and it is evident from the experiments of Maurizio (1965) with bees, cited

B I O C H E M I S T R Y OF SUGARS AND POLYSACCHARIDES

299

above, that di- and even trisaccharides can be absorbed intact. This conflicts with the suggestion that insects may absorb only monosaccharides (Gilmour, 1961), although, in view of the activities of digestive enzymes, the latter is doubtless the major normal process. High blood glucose and fructose levels are generally restricted to insects which feed directly on plant juices of high sugar content, such as bees and aphids. An exception to this is the build-up of glucose in overwintering pupae of Celerio euphorbiae (Heller and Mochnacka, 1951), but this is at least restricted to a non-feeding stage which need not maintain a gradient for absorption from the gut. The absence of trehalose from blowfly maggots (Phormia: Evans and Dethier, 1957; Culliphora: Dutrieu, 1961a) may be related to the fact that these can subsist principally on protein and need not absorb carbohydrates. The absence of trehalose from the larval feeding stage of the phytophagous C. euphorbiue, however (Mochnacka and Petryszyn, 1959), does at present seem anomalous. B . R E G U L A T I O N OF B L O O D S U G A R

Despite the implication from a passive absorption mechanism that insects’ blood sugar may be at the mercy of their environment and food supply, there is clear evidence of regulation, at least with respect to trehalose. The levels of trehalose found range widely, but for a given species and stage, there does appear to be a characteristic trehalose level which tends to be maintained despite changes in nutrition. The latter are reflected more strongly in the glycogen reserves. Thus, silkworms kept undernourished for 1 week had the same blood trehalose as controls, but half the normal glycogen (Bricteux-Gregoire et ul., 1965). Horie (1960, 1961) noted that during starvation of silkworms the fat body glycogen content fell rapidly and blood trehalose declined only after some hours when glycogen was already minimal. This was confirmed by Saito (1963), who further showed that when the hemolymph trehalose level in fifth instar silkworms was experimentally lowered by bleeding and injection of saline the initial level was restored, at the expense of fat body glycogen, within 3 h. When blood trehalose was experimentally elevated by injection of this sugar, it soon declined, establishing a new steady state soniewhat above the norm within 3 h. In diapausing saturniid silkmoth pupae subjected to integumentary injury, hemolymph trehalose temporarily rises after several days and returns toward its normal level (Wyatt, 1961b, 1963b). In the cecropia pupa during 6 months of diapause and subsequent adult development,

300

G . R. WYATT

while fat body glycogen and blood glycerol undergo extreme changes, blood trehalose is rather constant, first at a level characteristic of diapause, and then at a new, higher level established early in development (Fig. 1). Experiments with [14C]glucosehave shown that trehalose is

2001

-

f

150-

V x.

n

f - loo2350 E \ tn

E

%

40-\

20-

Treholose '\

-*-.

25" I

30

6' I

0-r

60 90 120 Time from en3 of spinning (days)

. 0

150

0

FIG. 1. Levels of trehalose, glycogen and glycerol in pupae of Hyulophoru cecropia during diapause and subsequent development of the adult. One month after spinning, pupae were placed at 25", then from these a group was placed at 6", and from the latter a group was returned to 25°C at the times shown. Samples of six pupae were taken for analysis. Trehalose was estimated as the total anthrone-reactive material in hemolymph; glycerol was estimated in hemolymph by periodate and chromotropic acid ; glycogen was extracted from the bled body with perchloric acid, precipitated with alcohol and determined with anthrone reagent. The broken lines are extrapolations based on previous work (cf. Fig. 9). (Wyatt, unpublished experiments.)

not inert in these systems, but is undergoing constant turnover (Saito, 1963; Wyatt, 1963b and unpublished). Another interesting demonstration of hemolymph trehalose regulation is the circadian rhythm in its level in crickets (GryZZus domesticus)

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301

described by Nowosielski and Patton (1964). Blood trehalose exhibits a peak at 05.00 hours, which does not appear to depend on rhythms in feeding activity.

Iv. BIOSYNTHESIS AND

U T I L I Z A T I O N OF S U G A R S

A . GLUCOSE

1. Phosphatases In vertebrate animals, the release of free glucose, as a product of glycogenolysis or gluconeogenesis, is a function of glucose-6-phosphatase, an enzyme specific for this substrate (though also catalysing certain transfer functions; Nordlie and Arion, 1964) found in the endoplasmic reticulum of liver. The only report of this enzyme in an insect appears to be the recent study by Terner et al. (1965). Because of their interest in correlating specific enzyme production with polytene chromosome morphology, these workers examined salivary glands of Sciaru coprophila larvae by both histochemical (light and electron microscope) and biochemical methods. An enzymic activity was demonstrated that was specific for glucose-6-P, was maximal at pH 6.5, was inhibited by zinc and certain other ions that also inhibit the mammalian enzyme, and was localized in dilated cisternae of the endoplasmic reticulum. The enzyme first appeared during the fourth instar, and was highly active in the prepupa and pupa, which correlates with the period of evident glycogen storage in the cytoplasm of this gland. However, its absence from homogenates of whole second and third instar larvae indicates that the animal cannot be dependent on glucose-6-phosphatase for production of blood glucose. Weak activity against glucose-6-phosphate (7% of the rate for trehalose-6-phosphate) is exhibited by partially purified trehalose-6phosphatase from adult Phormia (Friedman, 1960a, b). Crude extracts from larval cecropia fat body hydrolyse these two sugar esters at about the same relative rates, which suggests that the same enzyme may be responsible (Murphy and Wyatt, 1965). Lack of glucose-6-phosphatase activity is also suggested by the presence of appreciable concentrations of glucose-6-P in the hemolymph of larval Bombyx mori (Wyatt and Kalf, 1957; Wyatt et al., 1963). An enzyme designated glucose-1-phosphatase, apparently specific for the hydrolysis of hexose-1-phosphates (and also the unnatural substrate p-nitrophenyl phosphate) was discovered in larval Bombyx mori hemolymph by Faulkner (1955). The enzyme is "without effect on glucose-6-P, and is inhibited by fluoride, phosphate and arsenate and,

302

G . R . WYATT

weakly, by citrate; the pH optimum is 4.5. Faulkner suggested that this enzyme would intermediate the release of glucose from glycogen, combining the steps catalysed in mammals by phosphoglucomutase and glucose-6-phosphatase, but no experimental support for this role was presented, nor was the distribution of the enzyme in the insect examined. Curiously enough, there seems to have been no further work on this unusual enzyme. 2. Hexokinases The presence of hexokinases is implied by the ability of insect tissues, for example muscle (Sacktor, 1955), fat body (Clements, 1959; Shigematsu, 1960) and tissue culture cells (Grace and Brzostowski, 1966), to metabolize free glucose. Since the only established pathway for metabolism of trehalose involves its hydrolysis to glucose by trehalase, the next step in this case too would be phosphorylation by a kinase. Hexokinase activity has been demonstrated in flight muscle of the housefly (Chefurka, 1954) and the locust (Kerly and Leaback, 1957) and in silkworm fat body (Shigematsu, 1958) and mid-gut (It0 and Horie, 1959), but characterization of it was rather slight. The enzynie is non-specific ; that from locust muscle catalysed the phosphorylation of glucose, fructose, mannose and glucosamine, and was saturated by glucose at a substantially lower concentration than by fructose. These characteristics resemble those of the mammalian brain enzyme, but unlike the latter locust muscle hexokinase is located in the soluble rather than a particulate fraction of the cell. A more recent and detailed report (Ruiz-Amil, 1962) concerns the hexokinase of the honey bee; 75% of the activity in the adult bee was found in the thorax, thus the enzyme presumably derives chiefly from muscle. It is soluble and has been purified 20-fold by precipitation and chromatography. It is nonspecific and, as with the mammalian enzyme, both the K,,,and the V,,, for fructose are higher than the corresponding parameters for glucose, so that the relative phosphorylation rates for these two sugars would depend upon their concentration; the precise kinetic constants of the honey bee enzyme, however, are distinct from those of any previously described hexokinase. The locust and honey bee enzymes, like their mammalian counterparts, are inhibited by glucose-6-P, which may be important in regulation of glycolysis. B . U S E OF M O N O S A C C H A R I D E S O T H E R T H A N G L U C O S E

The utilization of various carbohydrates for nutrition has been assessed in many insects on the basis of such criteria as survival, growth,

B I O C H E M I S T R Y OF S U G A R S A N D P O L Y S A C C H A R I D E S

303

and deposition of glycogen. This work has been treated in other reviews (Trager, 1953; Dadd, 1963; House, 1964a). We shall here omit also the vast subject of digestive enzymes. It is interesting to note, however, that a line of tissue culture cells isolated from Antheraea pernyi, grown in the absence of normal digestive tissue, can use sucrose as well as glucose and fructose from the medium (Grace and Brzostowski, 1966). The nutritional results with monosaccharides fall in the following pattern for most insects: glucose and fructose are used readily, mannose with variable efficiency, while galactose is used to a relatively small extent, if at all. Among locusts, for example, growth in the early instars is normal on diets with glucose, fructose or mannose as the sole carbohydrate, but on mannose it fails in the fourth instar; galactose supports some growth in Schistocerca but none in Locusta (Dadd, 1960). Isolated fat body of Phormia regina converts glucose, fructose and mannose to trehalose with equal rapidity, while incorporation of galactose proceeds at about one-sixth the rate. After maintenance of Phormia adults on mannose injections for 6 days, the only sugars found by chromatography of hemolymph were glucose and trehalose (Clegg and Evans, 1961). The survival of Bombyx silkworms is promoted equally by glucose, fructose and mannose and much less well (but above controls) by galactose (Ito, 1960). Honey bees and some related Hymenoptera are unusual in that mannose and galactose are toxic (Staudenmeyer, 1939; Maurizio, 1965). Sorbose is apparently not used by insects. Utilization of pentoses is generally poor but, again, there is variation; survival of Bombyx is promoted by xylose (lto, 1960). The metabolism of different monosaccharides requires appropriate enzymes (Hollmann, 1964). In vertebrate animals, fructose requires a fructokinase which produces fructose-1-phosphate and enzymes to lead the latter into the glycolytic pathway; although hexokinase can phosphorylate fructose in vitro, this is inhibited in vivo because of the much greater affinity of the enzyme for glucose, and the characteristics of the insect enzyme, discussed above, would lead to the same expectation. Mannose can be effectively phosphorylated by hexokinase but a phosphomannose isomerase is required for its further metabolism. Galactose requires a specific kinase and other enzymes that catalyse its conversion to UDP-galactose and epimerization to the glucose analogue. The only report that I am aware of concerning such pathways in insects is that in which Sols et al. (1960) explain the enzymic basis of mannose toxicity in honey bees. Bees possess a high level of hexokinasC which phosphorylates mannose more efficiently than glucose, along with a negligible

304

G. R . WYATT

level of phosphomannose isomerase. Thus, mannose would competitively inhibit glucose phosphorylation and accumulate as mannose-6-P which, as a competitive inhibitor of phosphoglucose isomerase, would further interfere with glycolysis. C . B I O S Y N T H E S I S OF TREHALOSE

1. Synthesis in vivo It will be evident that the synthesis of trehalose must be a process of some importance in the economy of insects. Conversion of glucose to trehalose in vivo is rapid. When 0-14 mg of [14C]glucosewas injected into adult Schistocerca, 92% of the radioactivity in hemolymph collected 15 min later was in the form of trehalose; with a smaller injected dose, the percentage was still higher (Treherne, 1958~).In adult Phormia after injection of tracer amounts of [14C]glucose, the radioactivity on sugar chromatograms was 50% in trehalose at 2 min and 90% after 10 min (Clegg and Evans, 1961). Incorporation in the fifth-instar cecropia larva appears to be somewhat less rapid (Table II), but the TABLE I1 Incorporation of glucose into trehalose in cecropia silkworm larvae*

Expt.

Time (min)

1 2

10 30

Distribution of radioactivity in sugars (%) Glucose Trehalose

120

79 3.6 1 *7 1 .o

300

0.1

60

21 96-4

98.3 99.0 99.9

* [l-14C]glucose(2 pc in 3 pmoles) was injected into feeding late fifth-stage larvae. Hemolymph was collected, deproteinized, and deionized, the neutral solutes were separated on paper chromatograms, and glucose and trehalose were eluted and counted. Radioautographs showed activity only in these sugars. In Expt. 1 tissues were dissected from the larva at 10 min; in Expt. 2, successive bleedings were taken from a single larva. (G. R. Wyatt, unpublished experiments.) abundant hemolymph from which the glucose must be withdrawn has to be taken into account. After 10 min of incorporation (Table 11, Expt. l), the specitic activity of trehalose was much higher in the fat

BIOCHEMISTRY O F SUGARS A N D POLYSACCHARIDES

305

body (5,660 cts/min per mg) than in the hemolymph (1,490 ctslmin per mg), which demonstrates synthesis in this tissue. Because of uncertainty in the specific activity of the precursor glucose pool, absolute rates of trehalose production cannot be calculated in these experiments ; such rates must vary greatly with physiological conditions, and it would be of interest to determine them in vivo.

I I

I I

Treholose

G -1 - p

--

I Arnylose I I

1 I

HO ,’ Phospho-

I I 1

rylose (c)

I

I

I

I UDP

I

/ /

TREHALOSE

FIG. 2. Metabolic pathways linking glucose, trehalose and glycogen in insects. Regulatory mechanisms: (a) hexokinase is inhibited by glucose-6-phosphate; (b) glycogen synthetase is activated by glucose-6-phosphate; (c) glycogen phosphorylase is activated by AMP and under hormonal control; (d) trehalose phosphate synthetase is activated by glucose-dphosphate and inhibited by trehalose; (e) trehalase is inhibited in hemolymph by a protein-ion complex and in muscle by means unknown.

Incorporation of isotopefrom [14C]pyruvateand [14C]glucose-l-Pinto blood trehalose, fat body trehalose and fat body glycogen in the fifthinstar silkworm has been measured by Bricteux-Gregoire et al. (1964, 1965). The larvae were injected, then kept 4 h at 25°C without food, then bled and dissected. From both precursors, the specific activity of the glycogen was less than that of the trehalose in the fat body, and the latter in turn was lower than trehalose in the hernolymph. It was suggested that this shows that glycogen is not an obligatory intermediate in the biosynthesis of trehalose and that it may indicate a site of trehalose synthesis other than fat body (Florkin and Jeuniaux, 1965). One may inferfrom observations on the cecropia silkworm,however (Table II), that the precursor glucose pool would have very low radioactivity

306

G . R . WYATT

by 4 h, and that trehalose is made and discharged from the fat body so rapidly that after this length of time a low specific activity would be expected at the site of synthesis. This would be especially the case during starvation, since breakdown of reserve glycogen would be flushing out the intracellular sugar pool. Glucose-1-P was an efficient trehalose precursor in these experiments (15% of the injected activity was found in trehalose), although sugar phosphates are usually considered to enter cells poorly; but it is possible that this is attributable to the activity of hemolymph glucose-1-phosphatase (Faulkner, 1955). Net changes in blood trehalose in Antheraea pernyi larvae after injection of sugars have been measured by Egorova and Smolin (1962b). Glucose (40 mg) caused a doubling of blood trehalose at 4 h, followed by a fall to normal levels; glucose-1-P (90 mg) caused a slight rise, and sucrose (40 mg) had little or no effect.

2. Synthesis by tissues and homogenates The activity of the fat body in trehalose synthesis has been unequivocally shown by incubating fat body tissue in uitro with 14Cglucose and then isolating radioactive trehalose. Such experiments have been done by Clements (1959) and by Candy and Kilby (1959) with fat body from Schistocerca gregaria, by Clegg and Evans (1961) with Phormia regina and Leucophaea maderae, by Saito (1963) with Bombyx mori and by Murphy and Wyatt (1965) with Hyalophora cecropia. The efficiency of this conversion may be quite high: Clegg and Evans used L. maderae fat body to prepare [14C]trehalosewith specific activity 40-60% of that of the precursor glucose. When Phormia fat body was incubated without glucose, there was net release into the medium of trehalose which must have originated from endogenous glycogen (Clegg and Evans, 1961). Fat body, however, appears not to be the sole site of trehalose synthesis. Hines and Smith (1963) incubated radioactive glucose, with homogenates of several locust tissues, fortified with ATP, for 1 h, and found incorporation into chromatographically isolated trehalose in preparations from leg muscle and head tissues as well as fat body. Trivelloni (1960) also detected trehalose synthesis in locust muscle homogenate. Shyamala and Bhat (1965) recently demonstrated a small conversion of labelled glucose to trehalose by perfused isolated silkworm mid-gut, though this was much less than the conversion after absorption in uiuo. Clegg and Evans (1961), on the other hand, incubating isolated Phormia tissues with [14C]glucose,found no incorporation into trehalose by mid-gut, and a small amount, which they felt might be

B I O C H E M I S T R Y OF SUGARS A N D P O L Y S A C C H A R I D E S

307

due to contaminatingfat body, by flight muscle preparations. Considering the relative bulk of the fat body in most insects, its contribution to trehalose production in the intact animal must be by far predominant (despite a contrary report by Egorova and Smolin 1962~). 3. Enzymes of trehalose synthesis Synthesis of trehalose by an insect enzyme was first achieved by Zebe and McShan (1959), who incubated a 20% solution of glucose with Leucophaea muscle trehalase for many hours and demonstrated by chromatography that some trehalose was formed by the reversal of hydrolysis. With the levels of sugars that prevail in insects, however, this reaction would proceed solely in the degradative direction. The enzyme system specific for trehalose synthesis in insect fat body was first examined by Candy and Kilby (1961), and evidence was obtained in support of the reaction sequence previously discovered in yeast by Cabib and Leloir (1958): (1) Trehalose phosphate synthetase: UDP-glucose

+ glucose-tip

trehalose-6-P

-+

+ UDP

(2) Trehalose-6-phosphatase: Trehalose-6-P

+ H 2 0 + trehalose + Pi

Using an extract of adult Schistocerca fat body, Candy and Kilby showed that [14C]glucose incubated together with ATP and UDPglucose was incorporated rather efficiently into trehalose. Chromatography of reaction mixtures taken at different times showed that radioactivity appeared first in glucose-6-P,then in trehalose-6-P, and then in free trehalose, in accordance with the above sequence. In partially fractionated fat body extracts, the required accessory enzyme activities of phosphoglucomutase, nucleoside diphosphate kinase, UDP-glucose pyrophosphorylase and trehalose-6-phosphatase were demonstrated. Trehalose-6-phosphatase,a hydrolase specific for this substrate maximally active at pH 7, has been partially purified from Phormia regina by Friedman (1960b). Trehalose phosphate synthetase from larval fat body of Hyalophoru cecropia has recently been studied in some detail by Murphy and Wyatt (1965). The enzyme was obtained free of glycogen synthetase by highspeed centrifugation of fat body homogenates, during which the latter enzyme sedimented with the particulate glycogen. When the supernatant fraction was incubated with UDP-glucose and glucose-6-P, the release of UDP (assayed enzymically) was stoichiometric with the incorporation of glucose (measured by radioactivity) into trehalose.

308

G. R. W Y A T T

Trehalose-6-phosphate did not accumulate because the preparations contained highly active trehalose-6-phosphatase.The most interesting characteristic of the synthetase is that it is inhibited by trehalose and exhibits other features which are interpretable as allosteric effects. With dialysed enzyme preparations and no added Mg++,trehalose at 35 mM inhibited release of UDP from UDP-glucose by more than 90%, but the extent of inhibition varied not only with trehalose concentration but also with the levels of Mg+ and glucose-6-P (Fig. 3). Magnesium +

r

1

I

I

I

1

Trehalose (mM)

FIG.3. Inhibition of trehalose phosphate synthetase by trehalose. Fractions from Hyalophoru cecropia larval fat body homogenate were assayed for release of UDP from UDP-glucose in the presence of glucose-6-phosphate: X, 37,000 x g pellet, containing glycogen synthetase, not inhibited by trehalose; 0, 37,000 x g supernatant, containing trehalose phosphate synthetase, assayed without Mg+ ; 0 , same, assayed with 20 m~ Mg++.(From Murphy and Wyatt, 1965.) +

enhanced the binding of glucose-6-P, and the interaction between the latter compound and trehalose, although appearing to be competitive, is believed to involve separate sites on the enzyme. The presence of two or more interacting sites for glucose-6-P itself, as substrate, is implied by the sigmoid curve that is obtained when reaction velocity is plotted against its concentration. Both the inhibition by trehalose and the “co-operative” kinetics for glucose-6-P are lost when the enzyme is mildly maltreated (for example, by incubation at 43°C for 30 min, or addition of M mercuric acetate), although its catalytic activity is retained. Even precipitation by ammonium sulfate caused some loss

BIOCHEMISTRY OF SUGARS AND POLYSACCHARIDES

309

of the enzyme’s peculiarities, and this unfortunately prevented study of them with purified material. All these properties are closely analogous to those recently recognized in a number of enzymes which catalyse regulatory rate-limiting steps of metabolic reaction sequences in micro-organismsand animals. These properties have been discussed and designated by the term “allosteric” by Monod et al. (1963, 1965). It is suggested that inhibition of trehalose phosphate synthetase by trehalose may be important in the natural regulation of the rate of trehalose synthesis and, accordingly, in the homeostasis of hemolymph trehalose levels. It should be noted that this is not a simple product inhibition, and that the immediate product of the enzyme, trehalose-6-P, cannot accumulate because of the activity of the phosphatase. In vitro, the synthetase can be inhibited by trehalose-6-P (and by cellobiose), but this is not believed to have physiological sigmficance. Evidence for control of trehalose synthesis in fat body by the ambient trehalose level was sought in experiments with pieces of this tissue incubated in physiological media (Murphy and Wyatt, 1965). Addition of trehalose at 50 m ~which , is the approximate level in the hemolymph of mature cecropia larva, was shown to inhibit the incorporation of glucose into trehalose, and concomitantly to stimulate incorporation into glycogen. The degree of inhibition of trehalose synthesis, however, was variable (about 30% and 75% in two experiments reported) and seems insufficient to account for the homeostasis of hemolymph trehalose. Further study of environmental influences in fat body metabolism, and comparison of rates in vitro and in ziiz70, are needed. The question arises, in considering such a mechanism, how the characteristically different trehalose levels of different developmentalstages are established. As noted above, the sensitivity of the synthetase to inhibition by trehalose can be altered by the levels of Mg+ and glucose-6-P, and it is conceivable that these levels in fat body might be subject to hormonal or other influences. The activity of the synthetase might be modiiied by hormones in other ways as yet unknown. It should also be noted that the level of trehalose in the hemolymph is the resultant of its rates of synthesis and degradation. The latter process will be discussed next. +

D . CLEAVAGE A N D USE OF TREHALOSE

1. Cleavage of trehalose by insect tissues The metabolic use of trehalose depends upon its hydrolysis to glucose by the enzyme trehalase: Trehalose

+ Ha0

i -

2 glucose.

310

G . R. WYATT

No other mechanism for the metabolism of trehalose has been established. Trehalase in insects was first demonstrated, before the recognition of trehalose in them, by Frerejacque (1941). In seeking a source of this enzyme, he adopted the reasonable approach of examining insects that fed upon fungi, which were known to contain trehalose. Finding trehalase in these, he then tested other insect species and found, surprisingly, that a wide variety, regardless of diet, yielded active preparations. In the potato beetle, the head, thorax and abdomen all contained trehalase. Trehalose was split much more rapidly than sucrose or maltose. The pH optimum was 5.8. Activity was stated to be greater in phosphate than in phthalate buffer, which led to the suggestion that cleavage of trehalose might involve phosphorolysis. Some degree of enhancement of trehalase activity by phosphate has been confirmed with certain preparations from insect muscle (Kalf, 1957; Gilby et al., 1966), and the suggested phosphorolytic cleavage has been discussed (Kalf and Rieder, 1958). This would be analogous to bacterial sucrose phosphorylase, which converts sucrose directly to fructose and glucose-1-phosphate (Doudoroff et al., 1948). Highly purified soluble insect trehalase, however, and some well washed particulate preparations from muscle, show no effect of phosphate (Friedman, 1960a; Zebe and McShan, 1959). Furthermore, 2 moles of glucose have been obtained from each mole of trehalose with both types of preparation. It now appears that such effects of phosphate as have been observed are due to some sort of enzyme activation (Gilby et al., 1966), and it can be stated that there is no firm evidence for any nonhydrolytic cleavage of trehalose in insects. A number of workers have assayed different insect tissues for trehalase activity. Most of their observations have been qualitative or semi-quantitative, and in many cases only the digestive tract was examined (Table 111). Some experiments directed toward determining the quantitative distribution of trehalase in tissues of three species of Lepidoptera are summarized in Table IV. The digestive system exhibits high trehalase activity in every insect species in which it has been examined, and, while findings differ somewhat, the activity seems generally to be greatest in the salivary glands and mid-gut. Activity has also been found in muscle whenever it has been tested (with the exception of leg muscle from Melanoplus direrentialis; Derr and Randall, 1966). From several papers, one would judge the level of activity in muscle to be low, but since muscle trehalase is largely bound to cell structure and is less stable than the intestinal enzyme (see below),

BIOCHEMISTRY OF SUGARS A N D POLYSACCHARIDES

311

TABLE 111 Trehalase activity in insect tissues* Order and species Orthoptera BIaberus discoidalis

Leucophaea maderae

Melanoplus differentialis

Schistocerca gregaria Schistocerca gregaria

Homoptera Aphis fabae Aphids (4 other species) Megoura viciae Lepidoptera Bombyx rnori Chilo simplex Leucania separata Coleoptera Capnodis milliaris

Staget

Trehalase in tissues

++

Mid-gut and caeca ; Ehrhardt and Voss, 1962 other parts of gut, including salivary glands

+ + +, +, + +,

Fore-gut mid-gut Zebe and McShan, 1959 hind-gut ', muscle fat body f, hemolymph ' Malpighian tubules Den and Randall, 1966 fore-gut and mid-gut hind-gut &, thorleg acic muscle muscle, fat body and hemolymph hemolymph Howden and Kilby, 1956 Fat body

+ +,

+,

+,

++

+,

All parts of gut including Evans and Payne, 1964 salivary glands

+ Salivary glands + + ,crop +, mid- and hind-gut

++ ++

Duspiva, 1954

Salivary glands and gut Duspiva, 1954

+

Salivaryglands +,crop mid- and hind-gut

+,

Ehrhardt, 1962

++

+

Salivary glands Mukaiyama, 1961 Gut Yushima and Ishii, 1952 Hemolymph at moults Liu and Feng, 1965 only

+

+

Anterior, middle and pos- Courtois et al., 1964 terior parts of larva

++

Ips typographus

References

Whole insects

+

++

Courtois et al., 1961; Chararas et al., 1963 Allmann and-Duspiva, 1966 Chararas et al., 1963

Melolontha melolontha L, A Gut + L, P, A Whole insects Pissodes notatus Diptera Salivary glands Laufer et al., 1964 Chironomus thurnrni L Lucilia sericata L Salivary glands and mid- Evans and Marsden, 1956 gut Phormia regina A Flight muscle + Clegg and Evans, 1961

+

++ ++

+

* Degrees of trehalase activity are signified by are given in Tables IV and V. t Abbreviations as in Table I. 12-bA.I.P. 4

O,

+, + and + +. Additional

data

312

G. R. W Y A T T

TABLE IV Trehalase activity in tissues of some Lepidoptera. Celerio euphorbiae

Bombyx mori

(Petryszyn and Szarkowska, 1959)

(Saito,

pmoles T/5 h/100 rng powder

pmoles G/h/mg protein

mg G/30 min/100g fresh wt

pmoles G/min/mg protein3

L, V

L, mature Prepupa

L, V

1960)

(Duchiteau-Bosson et al., 1963)

Samia Cynthia ricini (Chang et al., 1964)

Activity in tissues Intestine Muscle Fat body Integument Silk gland Hemolyrnph

L, feeding?

P A

162

11 51

19 10 30

7 16 5 32

10

2

11.8 0-28 0.58 0

0

Fore- 7.5 Mid- 16.8 Hind- 11.8

8,068

0

138 7 0,225

2.3 1.1

0 0

* Petryszyn and Szarkowska precipitated the tissues with alcohol, extracted with water, and reprecipitated with alcohol, which may have led to some loss of activity. All other authors assayed activity directly in low-speed supernatants from tissue homogenates. Abbreviations as in Table I. t Prepupae gave similar distribution. 2 The units are translated from the Chinese as per minute, although this gives exceptionally high activities, and the results of Saito (per hour) are incorrectly cited by Chang et al. as per minute. 5 Two samples, consisting of tracheae and epidermis with minimal muscle contamination. part of it may often have been lost in preparing extracts for assay. Fat body from several insects has given low trehalase activity, but in Sumiu ricini it is reported to be sharply elevated at the time of the larval moult (Chang et ul., 1964). In examining epidermis, the problem is to obtain samples free of adhering muscle; DuchSiteau-Bosson et al. (1963)have attempted this by taking from silkworms the tracheal trunks, with their epidermal layer and adhering sheets of epidermis torn from the cuticle, and they believe that their results indicate absence of trehalase from the epidermis. Silk gland has given zero activity in three different laboratories. Data on hemolymph range from no trehalase to highly active, and the reasons for this interesting variation will be discussed below.

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2. Characteristics of insect trehalases Some properties of insect trehalase preparations are set forth in Table V. The enzyme has been substantially purified from whole insects of several species, and, in addition, particulate preparations from insect muscle have been studied. It has recently become evident that two distinct types of trehalase are represented, and that both types can be found in the same insect (Gussin and Wyatt, 1964, 1965; Gilby et al., 1966). Both are specific for trehalose and are not general a-glucosidases. The soluble trehalase which has been purified will be referred to as gut trehalase, since enzyme activity with similar characteristics can be demonstrated in gut extracts. The enzyme from muscle will be termed muscle trehalase. In the preparations purified from whole insects, muscle trehalase was probably eliminated during the fractionation or inactivated (the muscle enzyme, for example, is labile to heat (Gilby et al., 1966), which forms a step in one preparation procedure (Kalf and Rieder, 1958)). Insect gut trehalase, from six species belonging to three orders, is similar in having a pH optimum in the range 5-0-5.7. The K,,, is in most cases in the neighbourhood of 0-5-1.0 m~ (of the two values for Bombyx mori, that obtained with purified enzyme is probably the more reliable; the relatively high value for Melanoplus, however, may indicate a real species difference). The enzyme from Bombyx pupae yielded two components during chromatography on DEAE-cellulose, but these had the same kinetic constants (Saito, 1960). No specific activators of gut trehalase have been reported. The enzyme from Phormia is stated not to be inhibited by sucrose (20 mM, when trehalose was 6 m ~ ) , nor by glucose at twenty times the substrate concentration, but tris(hydroxymethy1)-aninomethane inhibits 20% at 0.025 M, and there exists a natural inhibitor in hemolymph, which will be discussed shortly (Friedman, 1960a). The enzyme from Melanoplus is strongly inhibited by trehalosamine, but not by glucosamine (Derr and Randall, 1966). A k s t estimate, by means of gel filtration, of the molecular weight of Blaberus discoidalis gut trehalase gave a value of about 70,000 (Gilby et al., 1966). The properties of muscle trehalase have been studied in particulate preparations obtained by fractional centrifugation of thoracic muscle homogenates. Both the pN optima and the K,,, values of the enzymes from several insect species fall in slightly higher ranges than those of the gut enzymes. For two species (Hyalophora cecropia and BZ. discoidalis), muscle and gut trehalases have been compared under identical conditions, and it is certain that the differences are real; for Phormia regina, the rather small difference between the constants measured in two

TABLE V Properties of insect trehalases Type of enzyme; species source

Gut type BIaberus discoidalis Melanoplus differentialis Bombyx rnori B. mori Galleria mellonella

PH optimum

K,(mM)

Mid-gut 100,OOOg supernatant, dialysed Purified 50 x from whole adults

5.0

0.5

5.5

5-1

Larval mid-gut 3,000 rev/min supernatant Purified 200 x from whole Pupae Purified 50 x from whole larvae

5.4

2.9

Preparation

5.2 5.5

2 P Remarks

References Gilby et al., 1966

E. = 7,160 cal/mole

Derr and Randall, 1966 Horie, 1959

0 4 - 0 . 4 7 Em = 9,500 cal/mole; two 1.3

Saito, 1960 chromatographic components Stated K , of 0.13 mM is erro- Kalf and Rider, 1958 neous; see Gussin and Wyatt, 1965

Hyalophora cecropia Phormia regina

Muscle type Bl. discoidalis

Leucophaea maderae H. cecropia Phormia regina

Mixed? Melolontha vulgaris

Larval mid-gut 100,OOO g supernatant Purified 1,OOO x from whole adults

5.7

Adult thoracic muscle microsomes

6.0

Thoracic muscle homogenate

6.0

Adult thoracic muscle microsomes Adult flight muscle mitochondria

6.5

3.6

5-8

1-3

Precipitated from supernatant from whole adults

6.5

0.7

0.4

Gussin and Wyatt, 1965

5.6

0.67

Friedman, 1960a

n grr

e *4

.e 4

3.3

Activated by various treatments; K,,, of activated enzyme = 1.7 mM; E. = 15,000 cal/mole Activated by freezing and thawing Activated by freezing and thawing, oleic acid Muscle soluble fraction has similar enzyme

Gilby et al., 1966

Zebe and McShan, 1959 Gussin and Wyatt, 1965 Hansen, 1966 Courtois et al., 1962

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laboratories could be due to differences in conditions, such as temperature, as Hansen (1966) has suggested. Muscle trehalase from cockroaches and H. cecropia is inhibited competitively by sucrose and more weakly by glucose and glucosamine (Zebe and McShan, 1959; Gussin and Wyatt, 1965; Gilby et al., 1966). The trehalase prepared from cockchafers (Melolontha) by Courtois et al. (1962), having a relatively high pH optimum and low value of K,, does not clearly fall into either of the classes of insect trehalase that have been discussed. Possibly the characteristics of the coleopteran enzymes are somewhat different, and possibly this preparation contained a proportion of both intestinal and muscle types. Trehalase preparations from several non-insect materials have kinetic characteristics quite like those of the insect gut enzymes: hog intestine (Dahlquist, 1960), Neurospora (Hill and Sussman, 1963), a cellular slime mould (Ceccarini, 1966). Yeasts, like insects, appear to yield at least two distinct trehalases: one with pH optimum 5-7 and K,,, 0-4 m~ (Panek and Souza, 1964), and one with pH optimum 6.9 and K, 10 m~ (Avigad et al., 1965). In insect muscle, the bulk of the trehalase is bound to cell structure, and its precise localization is an important question. After centrifugation of muscle homogenates at 100,OOO x g for 20 or 30 min (or longer), the fraction of total activity remaining in the supernatant was about 25% in P. regina (Hansen, 1966) and less in H. cecropia (Gussin and Wyatt, 1965) and Bl. discoidalis (Gilby et al., 1966). The soluble component from Phormia had the same kinetic constants as the particulate enzyme. In cecropia and Blaberus, centrifugation at 100,OOO x g yielded a microsomal fraction which contained half or more of the total enzyme with relatively high specific activity. Substantial activity remained with a low-speed fraction that contained myofibrils and nuclei, but mitochondria (isolated from cecropia only) showed very low activity. In Leucophaea maderae, on the other hand, purified mitochondria were reported to be highly active (Zebe and McShan, 1959); this has been confirmed for Phormia, in which it was also shown that fibrils are inactive, and it was inferred that the mitochondria are the chief seat of muscle trehalase. From the publication on Phormia (Hansen, 1966), it is not clear that a microsomal fraction was tested. The differences in cytologicallocalization of trehalase may be related to structural differences in the muscles of different insect groups, for the synchronous flight muscles of such insects as moths and cockroaches have a highlydeveloped sarcoplasmic reticulum, whereas the asynchronous muscle of flies has very little (Smith, 1961). But it would be worthwhile to seek

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G. R. W Y A T T

a common ground in these observations, recognizing the tendency of lipoprotein membranes to adhere to particulate cell components. It should also be pointed out that “microsome” is an operational term signifying a fraction obtained from tissue homogenates by highspeed centrifugation. It usually consists chiefly of fragments of endoplasmic reticulum, but may contain other structures, such as fragments of plasma membrane. In insect flight muscle, the plasma membrane undergoes deep tubular invaginations which make contact with the sarcoplasmic reticulum in structures known as dyads (Smith, 1961). This system is well developed in the cockroach, Periplaneta americana, and is believed to provide a pathway for the transmission of excitation to the interior of the fibre (O’Connor et al., 1965), but it seems worth considering that trehalose might enter by the same route. It would be of extreme interest to establish the precise localization of muscle trehalase by cytochemical means. A remarkable property of particulate trehalase preparations from insect thoracic muscle is that their enzyme activity is greatly increased as a result of freezing and thawing and certain other treatments which would tend to disrupt lipoprotein structure. The effect of freezing and thawing was first noted for L. maderae by Zebe and McShan (1959). Activation by this treatment, and also by oleic acid, was established for H. cecropia by Gussin and Wyatt (1965). The phenomenon has now been more fully studied with microsomal enzyme from B. discoidalis, and it is found that activity is enhanced five to tenfold by repeated freezing and thawing, by treatment with various detergents, or incubation with snake venom (Gilby et al., 1966). Snake venom retains power to activate trehalase when boiled, and this is believed to be due to its heat-stable phospholipase A. After such activation, the pH optimum of muscle trehalase is apparently the same as that of the original microsoma1 preparation but the K,,, is lowered to approximately one-half (1.7 mM). The activation is believed to involve an unmasking of enzyme embedded in lipoprotein membrane structure. Exhaustive snake venom treatment releases the enzyme as a homogeneous protein with molecular weight about 65,000-80,000, and deoxycholate also renders it nonsedimentable, but other activating treatments were less effective in solubilking. Curiously enough, the trehalase in homogenates of flight muscle from the flies Musca domestica,Phormia sericata and Sarcophnga bullata, although structure-bound, was not activated at all by freezing and thawing. The effects of other treatments with these species are not reported. The characteristics of trehalase known to originate from insect tissues other than intestine and muscle have not been examined.

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E. PHYSIOLOGICAL ROLES OF TREHALOSE A N D TREHALASE

1. Muscle Any tissue which is to degrade trehalose for the purposes of its internal energy supply requires trehalase, and this is clearly the function of the enzyme in muscle. Sacktor (1955) showed some years ago, before trehalose was recognized in insect hemolymph, that this sugar can support respiration in housefly flight muscle homogenates as readily as glucose or any other substrate. Clegg and Evans (1961), in a study of the physiology of trehalose and flight in adult Phormiu reginu, which we have already cited several times, showed that this sugar provides the major direct source of flight energy. During prolonged flight the blood trehalose level declines greatly and the wing-beat frequency declines in synchrony with it, while blood glucose remains almost unchanged (Fig. 4). Wing-beat frequency was restored equally

25r--------i

I-

01

Lp Glucose-

I

II

10

9

8

I

7

Decreasing WBF ( x IO3cycles/rnin)

Fro. 4. Relationship between the wing-beat frequency (WBF) and the concentrations of blood glucose and trehalose during prolonged flight of Phormia regina. Flies were flown until the desired WBF was reached, and samples of bload were then taken and assayed for sugars. Each point is averaged from at least ten individuals, and the vertical ban represent standard errors. (From Clegg and Evans, 1961.)

318

G. R. WYATT

after injections of trehalose or glucose, but because of the relative concentrations normally present in the blood, the former is considered to be quantitatively the more important metabolite. During prolonged flight, the major ultimate energy sources are calculated to be sugars stored in the crop and fat body glycogen; the former are transported to the flight muscle through the blood either directly or via conversion to trehalose in the fat body, while the latter is released for transport in the form of trehalose. Use of trehalose (as well as glycogen) during flight has also been demonstrated in Locusta migratoria (Bucher and Klingenberg, 1958) and in the cockroach, Periplaneta americana (Polacek and Kubista, 1960). From the simple facts that blood trehalose level falls during flight but is maintained during rest, and that the use of trehalose depends on specific, irreversible hydrolysis by trehalase, it follows that the activity of this enzyme must be regulated in relation to flight muscle activity. This is substantiated by recent analytical determinations of the levels of various substrates and metabolic intermediates in the thoraces of Phormiu regina taken after different durations of flight (Sacktor and Wormser-Shavit, 1966). Trehalose fell rapidly during the first minute of flight, and steadily thereafter (Fig. 5). Glucose, on the other hand 8,

I

I

3 2 r

I

r

I

0 1 2

Glycogen

0

15

I

45

30

60

0.10

GIu-6-P

o'20 0.15

0

15

30

45

60 0 Minutes

15

30

45

60

FIG.5. Concentrations of certain carbohydrates in thoraces of blowflies (Phormiu reginu) during 1 h of flight. Insets with expanded time scale show alterations at the initiation of flight. Each point is the average of three to twelve extracts, each made from ten flies. (From Sacktor and Wormser-Shavit, 1966.)

rose at the shortest time tested (5 sec), then fell and regained a steady state. Since the decline in glycogen was delayed relative to that of trehalose, and glucose-6-phosphate (a product of glycogen) did not

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rise at short times, the rise in glucose must be attributed to cleavage of trehalose rather than glycogen. The extreme rapidity of the rise in glucose indicates a control mechanism connected rather directly with the initiation of muscle contraction; of the many other metabolites measured, only AMP showed maximal change as early as 5 sec, and the peak in fructose diphosphate,which presumably results from change in AMP and ATP, was at 15 sec of flight. The mechanism of this trehalase activation is quite unknown. The experimentally demonstrable activation of muscle trehalase by freezing and thawing and other treatments discussed above at first seemed likely to have some relation to a metabolic regulatory mechanism, but the fact that trehalase in the muscle of flies is unaffected by freezing and thawing casts doubt on this speculation (Gussin and Wyatt, 1965; Gilby et al., 1966). The theoretical aspect of fuel and oxygen supply for insect muscle has been considered by Weis-Fogh (1964). It was assumed, with some prescience, that “trehalose in the blood is split to glucose on its passage through the fibre membrane.” Transport into the interior of the muscles and fibres is achieved by diffusion and muscular pumping, the latter being an inefficient tidal process resulting from the movement of blood into and out of intramuscular spaces as the muscle expands and contracts. Because of the long distances for diffusion and the small volume of the internal spaces, it is calculated that very high concentrations (of the order of 0-5-1%) of trehalose or other fuel in the hemolymph will be required in order to provide gradients that can supply the internal sites of metabolism at the rates required for the exceptionally high activity of this tissue. It is suggested that “the very high concentration of trehalose in insect blood can be considered a direct and essential adaption to flight.” The limiting dependence of muscle energy supply upon trehalose concentration is demonstrated by Clegg and Evans’ observation that blowfly wing-beat rate falls as blood trehalose falls. 2. Intestine The universal presence of high trehalase activity in the insect gut seems at first enigmatic, since trehalose is only rarely an appreciable component of the insect dietary. Although, as first shown by Philips (1927) for bees, insects can efficiently utilize ingested trehalose, it is clear that food digestion is not the normal function of gut trehalase. This was first remarked by Duspiva (1954), as a consequence of his finding (before trehalose was recognized in insects) that the salivary glands and intestines of several species of aphids exhibited much greater activity against trehalose than against sucrose, their most 12f

320

G. R. WYATT

abundant dietary sugar. He questioned tentatively whether symbionts might be involved. Petryszyn and Szarkowska (1959), after demonstrating high trehalase activity in the gut of Celerio euphorbiae, suggested that this might be a non-specific enzyme active also on some polysaccharides present in the Euphorbia plant. As it has become clear that intestinal trehalase is endogenous to the insect and specific for trehalose, it has been stated that this enzyme must play “un tout autre role que la digestion” (Chararas et al., 1963), but there has been no convincing proposal as to just what this role might be. Chang et al. (1964), noting that Samia ricini intestinal trehalase decreases in activity at the cessation of larval feeding, suggest that it might function in the synthesis of trehalose, but this is thermodynamically impossible. One may point out also that gut trehalase with its low pH optimum would be quite inactive under the alkaline conditions that prevail in the mid-gut of most Lepidoptera and some other insects (Waterhouse and Day, 1953). Thus, in silkworms in which the pH of mid-gut juice is 8-10, Horie (1959) estimated the pH optimum of mid-gut trehalase as 5-4 (with activity approaching zero at pH 85), while that of sucrase was 6.4 (activity more than half-maximal at pH 8), and that of amylase was 9.2. Horie also found trehalase only in the gut wall, and not in the digestive juice, but surprisingly he found the same thing for other intestinal enzymes (with the exception of amylase). The important question whether trehalase is secreted into the gut lumen has been examined in very few other insects; in Leucophaea maderae activity was found in both gut wall and contents, but in the fore-gut the ratio of trehalase to sucrase was much higher within the tissues (Zebe and McShan, 1959). Trehalase is present in the secretion from the salivary glands of Chironomus thummi (Laufer et al., 1963). The presence of trehalase in the insect intestine may be interpreted as a necessary corollary to the mode of sugar absorption in insects, to prevent loss of trehalose by diffusion. It has been shown that insects absorb non-specifically by diffusion through the gut wall, a gradient being maintained by conversion of the absorbed sugar to trehalose. Although monosaccharides are absorbed most readily, disaccharides and even in some instances trisaccharides pass the gut wall. Diffusion being freely reversible, it follows that there will be a tendency for trehalose, which is generally at high levels in the hemolymph, to escape through the gut wall into the lumen. The concentration gradient would prevent its re-absorption in the form of trehalose, but it will be reabsorbed if hydrolysed to glucose, the gradient for which is in the other direction, According to this hypothesis, then, the role of gut trehalase

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Is to prevent loss of trehalose in the excreta. It could do this most efficiently by splitting trehalose within the gut wall, the pH of which will be within the range of activity of the enzyme, but trehalase in the lumen could provide a second line of defense. This hypothesis implies a cycle, glucose being converted to trehalose in the fat body, and trehalose to glucose in the gut (Fig. 6). There is Epidermis, silkglond, etc

0 xidotion, biosynthesi

FIG.6. Interrelations of insect tissues in the metabolism of glucose, trehalose and glycogen. Conversions are indicated in their principal sites, and the scheme is simplified by the omission of apparently minor sites; for example, the synthesis of trehalose in tissues other than fat body, and the cleavage of trehalose in tissues other than muscle and intestine. G, Glucose; T, trehalose. For steps of intermediary metabolism, see Fig. 2.

some evidence for such a process. Clegg and Evans (1961) found by injection of [14C]trehaloseinto blowflies that it was converted to glucose within the intact insect at a rather substantial rate (an average of 20% in 15 min), whereas no such conversion took place in hemolymph in vitro. They suggest that the cleavage in vivo was caused by muscle trehalase, but it could equally well have been a function of the gut, for it seems likely that the trehalase ofintact resting muscle is somehow kept in check. That trehalose can escape via the gut in some instances is shown by its occasional presence in honeydew (Ehrhardt, 1962)and its presence in manna which is the dried excretion of scale insects (Leibowitz, 1943). The proposed cycle involves some use of energy for continual resynthesis of trehalose, but this is much less than the loss would be if trehalose were continually to escape. The rate of the cycle is presumably limited by the permeability of the gut cells to trehalose, which must be much less than their permeability to glucose. On the ledger against the

322

G. R . W Y A T T

energetic expense of the cycle is the saving that results from the absence of active transport mechanisms for sugars, which would otherwise be essential for absorption. Another aspect of the process is that it ensures a certain level of glucose in the hemolymph even when no carbohydrate has been ingested. This would be a substitute for the mammalian mechanism via glucose-6-phosphatase and would be important for the supply of carbohydrate to tissues that lack trehalase and cannot use trehalose directly. Ehrhardt (1962), although entertaining the possibility that trehalose in honeydew might originate from the hemolymph, thinks it more likely to be a product of transglucosylation in the gut, as are certain other oligosaccharides. But several attempts to demonstrate transglucosylation by trehalase from different sources have met with failure, as is indeed to be expected in view of the rigid specificity of this enzyme for both halves of the substrate molecule (Dahlquist, 1960; Courtois et al., 1962; Evans and Payne, 1964; Derr and Randall, 1966).

3. Hemolymph Although one would scarcely expect to find trehalase in the hemolymph, with its high content of trehalose, it is established that the enzyme is sometimes present. This was first reported for locusts in a brief note by Howden and Kilby (1956). Friedman (1960a, b) then made the interesting observation that whereas the trehalose was quite stable when undiluted blood from adult Phormia regina was incubated, it was split to glucose at a substantial rate if the blood was first diluted 100-fold with water. This was taken to indicate the presence of trehalase together with a dissociable inhibitor in the hemolymph and, in further studies (Friedman, 1961), such an inhibitor was demonstrated and to some extent characterized. When Phormia blood was heated to 60°C for 1-3 min (during which the trehalase is stable), and then incubated, trehalase was found to be active without dilution. That this depended upon inactivation of an inhibitor was established by adding untreated and heated blood to trehalase purified from whole insects. The trehalase in hemolymph also became active when EDTA (or, less effectively, citrate or oxalate) was added, which indicates a requirement for a divalent metal ion in the system. During dialysis, inhibitory effectiveness was somewhat lowered, and it could be restored by addition of Mg++.Thus, the system appeared to involve a heat-labile macromolecule, presumably protein, and a metal ion. Changes in the activity of hemolymph trehalase in relation to the moulting cycle were observed in Bombyx mori by Duchiiteau-Bosson

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et ul. (1963). They state that activity was found only during the moults, at which time the blood trehalose level was shown to fall sharply and elevated blood glucose had previously been observed (Florkin, 1937; Wyatt et al., 1956), but quantitative data on the enzyme were not given. These changes are interpretated in terms of a physiological de-inhibition of hemolymph trehalase, so that glucose is made available when needed for synthesis by tissues that lack trehalase, for example chitin by the epidermis and silk by the silkgland (Florkin and Jeuniaux, 1965). Quantitative data consistent with such a hypothesis have recently appeared in two papers from Peking (Chang et al., 1964; Liu and Feng, 1965). Chang et ul. have shown that in the eri-silkworm (Sumiu Cynthia ricini), hemolymph assayed directly shows trehalase activity during the moult and at the time of pupation, but not during the feeding stages. If treated with heat or EDTA (as shown by Friedman to de-inhibit trehalase in Phorrniu blood), activity is elevated and appears at stages when it was otherwise not detectable (Fig. 7b). The same treatments

M

P

FIG.7. Changes in trehalose and trehalase during fifth instar of the eri-silkworm (Sarnia Cynthia rieini; from Chang et al., 1964). M, Fourth moult; P, pupation. (a) Hemolymph trehalose content, determined with anthrone after treatment with alcohol, acid and alkali. (b) Activity of trehalase in hemolymph, assayed directly and after treatments that inactivate inhibitor. 0 , Untreated (but diluted 30-fold in the assay mixture); 0, heated 3 min a t 60°C before assay; A, treated with EDTA, final concentration 6 mM in phosphate buffer, before assay.

324

0 . R . WYATT

applied to gut and fat body homogenates did not enhance their trchalase activity. The blood trehalose level changes during development in a pattern approximately reciprocal to that of the enzyme (Fig. 7a). During starvation of larvae blood trehalase was also found to rise as trehalose fell. Liu and Feng (1965) showed rather similar changes during development in the armyworm, Leucania separata. Thus, trehalase in the hemolymph appears to have the role of making glucose available for synthesis by tissues that cannot use trehalose directly. This would presumably supplement the glucose that arises by cycling of trehalose through the gut, as proposed above. In considering the relation between these two processes, it is interesting to note that blood trehalase becomes active at times when normal gut activities are decreased as a result of cessation of food intake. The increased activity appears to result, in part, from the lifting of a natural inhibitor-an interesting phenomenon that requires further study. But is is evident (Fig. 7b) that the total level of trehalase in the blood also changes greatly. Biochemical characterization of the trehalase in hemolymph has not been reported, but it is presumed to be of the soluble, low-pH type (Friedman, 1960a, b), and its probable source is the intestine. F. DORMANCY A N D THE PROPERTIES OF TRBHALOSE

It has been pointed out by Clegg and Filosa (1961) that trehalose is found in particularly large amounts in the dormant stages of several diverse organisms. Thus, in the mould Neurospora tetrasperma, trehalose is present in dormant ascospores up to 14% of dry weight (about 50% of total carbohydrates), and after their activation trehalose is used rapidly and preferentially over other carbohydrates and lipids (Sussman and Lingappa, 1959; Sussman, 1961). The consumption of trehalose is correlated with some rise in the activity of trehalase (Hill and Sussman, 1964). Spores of the cellular slime mould, Dictyostelium mucoroides, contain more than 7% of their dry weight in trehalose, and this is used during germination and becomes undetectable in other lifecycle stages (Clegg and Filosa, 1961). In Ascaris Zumbricoides, which contains trehalose at all life stages, this sugar is found at especially high levels (7-8% of fresh weight) in the egg, but in this case the trehalose is largely not in the embryo but in the perivitelline fluid, and the possibility of its exercising an osmotic role in hatching has been suggested (Fairbairn and Passey, 1957). The cysts of the brine shrimp, Artemia salina, which are extraordinarily resistant to storage and desiccation, contain trehalose as some 15% of their dry weight (Dutrieu, 1959); this trehalose

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has been shown to be produced within the embryo itself and to accumulate only in those destined for dormancy. When these develop most of their trehalose is converted to glycogen and glycerol, the latter of which is believed to assist hatching by osmotic pressure (Clegg, 1964, 1965). In addition, Clegg and Filosa (1961) mention without data the finding of large amounts of trehalose in dried statoblasts of the bryozoan, Plwnatella, and the desiccatable eggs of the mosquito, Aedes aegypti. Among insects, accumulation of trehalose is not generally well correlated with the state of dormancy or diapause. Thus, in eggs of Bornbyx mori, although hibernated eggs contain more trehalose than nondiapause eggs, the level (4 mg/g) is not exceptionally high, and it increases during embryonic development in both types (Yamashita, 1965). In Hyalophora cecropia and other saturniids, the trehalose level is lower in the pupa in diapause than in either the mature larva or the developing adult (Table I; Fig. 1). But the principal hazard to which diapausing insects are exposed is not desiccation but freezing, and some protection against this is provided in many species by conversion of most of the carbohydrate reserves to glycerol and sorbitol (discussed below). In some other instances, as mentioned in an earlier section, accumulation of trehalose probably does play a role in conferring resistance to cold. Trehalose has several properties of potential biological value. It is exceptionally stable chemically, being subject neither to easy oxidation and degradation by mild alkali, like glucose, nor to hydrolysis by mild heat and acid, like sucrose. It is highly soluble, and unlike glycogen, osmotically active. It is converted by catalysed hydrolysis into glucose and thus its entry into the main pathways of carbohydrate metabolism can be regulated by control of the enzyme responsible for this one step. It is clear that among insects and other organisms, there is immense diversity of adaptation, and that the virtues of trehalose are made use of in relation to the needs of the organism in several ways. V . GLYCOGEN A . GLYCOGEN I N INSECTS

Glycogen has been recognized as an important constituent of insect tissues ever since the oft-quoted observations of Claude Bernard (1879) concerning its abundance in maggots. The extensive earlier work on the subject, which chiefly concerned the content ofnglycogen in insects, changes during metamorphosis, some nutritional aspects and presumed

326

G . R . WYATT

interconversions, was concisely reviewed by Babers (1 941), and some additional references may be found in the review by Rockstein (1950). One may remark at the outset that insect glycogen does not seem to have been examined for chain length and other structural characteristics, but it has been found to serve identically with commercial (presumably oyster) glycogen as a substrate for phosphorylase (Stevenson and Wyatt, 1964), and there is no reason to suspect any important chemical peculiarities in it. Among the more recent work, the distribution of glycogen in tissues of the larval and pupal blowfly, Phormia regina, has been meticulously examined in a histological study by Stay (1959). Almost all tissues (with the exception of the oenocytes, tracheal gland cells and parts of the digestive tract and ring gland) contain glycogen deposits, and these build up conspicuously toward the end of larval life. The greatest glycogen deposits in insects are commonly found in the fat body. Jn Bombyx mori, for example, during the fifth larval instar the glycogen content of the fat body rises from 5 to 20% of dry weight (Shigematsu and Takeshita, 1959). The tracheal organ of the Gastrophilus larva, a group of specialized cells associated with the fat body, attains 40% of its dry weight in glycogen (Levenbook, 1951). In a tissue such as fat body most of the glycogen is readily extractable; thus, Saito (1963) found that four extractions with cold trichloroacetic acid yielded 9.5% of the fresh weight of fifth-instar silkworm larval fat body as glycogen, and only a further 0.18% was obtained by digestion of the residue with hot 30% KOH. When a similar fractionation was applied to muscle, which contains much lower levels of glycogen, a much higher proportion of it was found to resist extraction (Kubista and Bartos, 1960). The glycogen extractable with cold acid was described as “free” and that not so extractable as “fixed”, and the contents of the two types in several preparations were : Locusta migratoria pterothorax, 0.23% and 0.20%; L . migratoria femur, 0.07% and 0.09%; Periplaneta americana metathorax, I -05% and 0.26%, respectively. Thus, the fixed glycogen was relatively constant while the free glycogen varied greatly in different tissues. A similar distribution in mammalian tissues, and a metabolic difference between the two fractions, has been described (Stetten et al., 1958). But it should be noted that the “fixed” glycogen from insect sources has not been chemically identified as such, and may include other protein-bound polysaccharides. The question of chemical identity must be kept in mind also when considering the records of glycogen in hemolymph (Table I). Glycogen has never been isolated and characterized from hemolymph, and while

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it is quite likely that there may be some glycogen free in hemolymph, particularly at times of histolysis, it is also likely that other polysaccharides from glycoproteins have been measured as glycogen. B. ACCUMULATION A N D CONVERSION D U R I N G GROWTH A N D METAMORPHOSIS

The problem of energy supply for metamorphosis, and the associated accumulation and conversions of nutrient reserves, form a classic subject for investigation. The early work on chemical changes during metamorphosis, including that on glycogen reserves, was summarized and discussed by Needham (I 929, 193l), and some subsequent papers are cited in the reviews mentioned above. Among the more recent papers, data on embryonic development in Bombyx mori are given by Chino (1957a, b) and Moulinier (1957) and in Tenebrio molitor by Ludwig and Ramazzotto (1965). Analyses on glycogen and other reserves during portions of post-embryonic development are recorded for the pea aphid, Acyrthosiphon pisum, by Srivastava (1965), the silkworms, B. mori (Zaluska, 1959), Antheraea pernyi (Smolin and Gudalina, 1957; Pravdina and Smolin, 1958), Samia Cynthia (Chang et al., 1963) and Hyalophora cecropia (Bade and Wyatt, 1962), the mealworm, T. molitor (Rousell, 1955), the dermestids, Anthrenus vorax (George and Nair, 1964) and A . grandis (Nettles and Betz, 1965), and the housefly, Musca domestica (Ludwig et al., 1964). The most usual pattern found from these analyses is a substantial rise in glycogen,both per individual and as percentage of body weight, during larval growth, interrupted (in those few cases where.this was examined) by a temporary decline at each moult (Fig. 8). In the fmal larval instar, the accumulation of glycogen is intensified in preparation for metamorphosis, and then during the pupal stage (in Holometabola) most of this is consumed as the chief energy source for development of the adult. The provocative observation was made for Bombyx mori as early as 1892 (Bataillon and Couvreur) that the glycogen content continues to rise for several days after the cessation of larval feeding, so that it must have some internal source. Since fat was found to decline in amount at the same time, conversion of fat to glycogen was proposed (Couvreur, 1895). The observations were confirmed for B. mori and similar changes at this particular stage of development have been found for several other insect species. The suggestion that carbohydrate is synthesized at the expense of fat was made by most of the earlier authors (see Bade and Wyatt, 1962), and some recent ones

328

0 . R. WYATT

(e.g. Ludwig et al., 1964; George and Nair, 1964). Since the net synthesis of carbohydrate from fat apparently cannot take place in higher animals (in contrast to the familiar reverse process) because of the irreversibility of the pyruvate oxidase reaction (Deuel and Morehouse, 1946), the possibility of this conversion in insects, if proven, would be of great interest. Most workers have not considered chitin among the carbohydrate reserves, however (nor trehalose, but this is quantitatively less).

I Moulting I

period Time (doys)

FIG. Changes in glycogen and chitin during the fourth larvs instar and moult in the silkworm, Bombyx mori. Glycogen: 4 -, males; -xfemales. Chitin: ---0---, males; ---x---, females. (From Zaluska, 1959.)

Zaluska (1959) has shown, by analyses at frequent intervals, that the chitin content of silkworms drops greatly in the prepupal stage, and the change is largely reciprocal with the rise in glycogen. This was also shown for the cecropia silkworm by Bade and Wyatt (1962). These authors suggest that chitin from the resorbed larval cuticle, possibly supplemented to some extent by protein and mucopolysaccharides, provides the material for the glycogen that is made after cessation of feeding. It is then unnecessary to invoke conversion from fat. The

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activity of one possible pathway for conversion of fat to carbohydrate, the glyoxylate cycle, was also sought but not found (Bade, 1962). It should be noted that Carpenter and Jaworski (1962) have found low levels of one of the enzymes of this pathway, isocitrate lyase, in prepupal Prodenia eridania, but the actual operation of the cycle, which they suggest, has not been demonstrated in an insect. It thus appears that the synthesis of carbohydrate from fat has not been demonstrated, and probably does not occur, in insects. This contrasts with the situation in Ascaris, in which such a process clearly takes place (Passey and Fairbairn, 1957). C . GLYCOGEN I N INSECT FLIGHT

In a recent review on the metabolism of insect muscle, Sacktor (1964) has fully discussed the chemical sources of muscular energy, and there is little to add. He points out that the distinction between insect groups which depend on fat for flight energy (Orthoptera, Homoptera and Lepidoptera) and those which consume carbohydrate is not absolute, for it is clear that some carbohydrate is burned by locusts, cockroaches and aphids. That the same is true for Lepidoptera is indicated by the recent demonstration of moderate levels of trehalase (Gussin and Wyatt, 1965)and glycogen phosphorylase (G. R. Wyatt and S. S. Wyatt, unpublished; see below) in thoracic muscle from the cecropia silkmoth, a species which is active in oxidizing lipids (Domroese and Gilbert, 1964). In insects in which carbohydrate is the major fuel, such as flies, it now appears that trehalose is a more immediate and critically regulated energy source than glycogen, as discussed above; nevertheless the numerous data showing depletion of glycogen during flight remain valid, and this is presumably brought about by some mechanism for activation of phosphorylase. A question which there has as yet been no attempt to answer is how the mobilization of glycogen reserves in tissues other than the muscle is initiated during prolonged flight. D . METABOLISM OF GLYCOGEN

1. Glycogen metabolism in fat body

Fat body glycogen constitutes a reserve that can be built up and drawn on as required. When insects are starved, this reserve is drawn on to maintain blood trehalose (see Section I11 B). During moulting, glycogen falls sharply, presumably because of the cessation of feeding and the demands of chitin synthesis (Zaluska, 1959). With the restora-

330

G . R. WYATT

tion of food intake, fat body glycogen is rapidly built up again. The functions of glycogen in fat body have been reviewed by Kilby (1965). There have been a few attempts to measure metabolism in intact fat body with radioisotopes. Clements (1959) incubated Schistocerca greguriu fat body in saline with [14C]glucosefor 4 h and, finding that only 3% of the total recovered radioactivity was in glycogen while 33% was in trehalose, concluded that glycogen appears to be only a minor product of fat body synthesis. Shigematsu (1960), however, in similar experiments with silkworm fat body, found about five times as much radioactivity in glycogen after incubation for 1 h as at 4 h. One may infer that (a) in such experiments it is essential to observe the kinetics of incorporation, and (b) in fat body in vitro the balance of metabolism is different from that in the intact animal so that glycogen is probably breaking down to release trehalose. The insect fat body is excellently suited for such studies, which might be directed to determine the nature of the controls over its metabolism.

2. Glycogen synthetase Despite the universality of glycogen in insects, and its profound involvement in their development and activity, very little study has been devoted to the enzymes of its metabolism. In vertebrate animals, it is now accepted that synthesisof the a-1,4 glycosidic bonds in glycogen is a function of UDP-glucose-glycogen transglycosylase (glycogen synthetase):

+

UDP-glucose glycogen,

-+

+

UDP glycogen, +

The enzyme in extracts from mammalian tissues is activated by glucose6-phosphate. Further analysis has shown that it exists in mammalian muscle and liver in two forms: the D form, which is dependent on glucose-6-P for activity and the I form which is independent (RosellPerez and Larner, 1964a). The D and I forms are interconvertible by mechanisms involving phosphorylation of the protein, and the proportions of the two found in a tissue varies in response to hormonal and other physiological conditions (Rosell-Perez and Larner, 1964b; Danforth, 1965). Glycogen synthetase was demonstrated in homogenates of locust muscle by incorporation of [14C]glucosefrom UDP-glucose into glycogen and by release of UDP from this substrate (Trivelloni, 1960), and it was also revealed in sections of the same material by a histochemical technique (Hess and Pearse, 1961). In the latter work, more activity was detected in leg muscle than in flight muscle, in contrast to the distribution of phosphorylase. Glycogen synthetase of fat body was

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examined by Vardanis (1963) with tissue from Periplaneta americana. The enzyme was demonstrated by incorporation from UDP-14Cglucose into glycogen and, in addition, the enzyme of UDP-glucose synthesis (UDP-glucose pyrophosphorylase) was detected by a spectrophotometric assay. In all three of these studies, glucose-6-P was tested as a potential activator, and found not to activate but even to inhibit slightly, and this was attributed to its entering into trehalose-P synthesis, which could compete for UDP-glucose. In the fat body extract used by Vardanis, the addition of glucose-1-P greatly increased the accumulation of radioactivity in glycogen and it was suggested that this phosphate might assume the role of glycogen synthetase activator in the insect. These preparations, however, contained glycogen phosphorylase activity at levels much higher than those of the synthetase, and it seems possible that the effect of glucose-1-P could be to shift the phosphorylase equilibrium, and so prevent loss of radioactivity from glycogen by phosphorolysis. In homogenates of fat body from larval cecropia silkworms, it was found that glycogen synthetase could be obtained largely free of trehalose-P synthetase and phosphorylase by high-speed centrifugation (Murphy and Wyatt, 1965). As in mammalian tissues, the synthetase was recovered bound to the pellet of particulate glycogen. In such preparations, activity measured by incorporation of radioactivity and by release of UDP from UDP-glucose were stoichiometrically approximately equal, and activity was strongly stimulated by glucose-6-P. Glucose-1-P was without effect, but galactose-6-P and glucosamine-6-P were stimulatory, which is in accord with the activator specificity of the mammalian enzymes (Rosell-Perez and Larner, 1964~).In most preparations, there was total dependence on glucose-6-P activation, but some late fifth instar larvae which had ceased feeding showed up to 35% of the activity in glucose-6-P independent form. This suggests physiological interconversion of two forms of the enzyme, as in mammals. Thus it appears that glucose-6-P is an activator of glycogen synthetase in insects, but different sensitivity to activation in different species, developmental stages and tissues is very likely. Such variations may be important in the control of glycogen deposition.

3. Glycogen phosphorylase This enzyme catalyses the reaction : Glycogen,, + phosphate

-

glycogen, -

+ glucose-1-P

332

G . R. WYATT

Since the action on glycogen amounts to removal or addition of glucose units at the ends of the branches of the macromolecule, the number of glycogen molecules is unchanged and glycogen can be eliminated from the equilibrium expression for the reaction. The latter then reduces to

the value of which at pH 7 is 3.1 manes and Maskell, 1942). From this it follows that in the test tube the reaction is readily reversible and proceeds easily in the direction of glycogen synthesis. Indeed, the enzyme is usually assayed by measuring release of inorganic phosphate from glucose-1-P in the presence of glycogen. In living cells, however, the ratio of inorganic phosphate to glucose-1-P is generally, if not always, much greater than 3, and under such conditionsthe phosphorylase reaction must proceed in the direction of glycogen breakdown. It is also observed both in vertebrates and in insects that changes in phosphorylase activity in tissues are correlated with rates of glycogenolysis, not with rates of synthesis, in vivo. In mammals and other vertebrates, the phosphorylases of muscle and liver are distinct enzymes, but each can exist in two forms: active and inactive. These are interconvertible by phosphorylation and dephosphorylation of the enzyme protein by a kinase (with ATP) and a phosphatase. The kinase is activated in liver by cyclic 3‘,5’-AMP, which is produced under hormonal influence, and thus glycogenolysis may be stimulated. In muscle, conversion of phosphorylase from its inactive form (phosphorylase b) to its active form (phosphorylase a ) occurs both by hormone action and by a mechanism connected with muscle contraction. In addition, phosphorylase b can be activated in a more direct sense by the presence of 5’-AMP, a nucleotide which would be formed by ATP cleavage in muscle contraction. Therefore, several levels of control exist. Knowledge of insect glycogen phosphorylases is fragmentary. Activity was demonstrated in silkworm (Bornbyx rnori) embryos (Shigematsu, 1956a, d), fat body (Shigematsu, 1956b) and mid-gut (It0 and Horie, 1959). Shigematsu assayed the enzyme in both directions of reaction, isolated crystalline glucose-1-P as a product from glycogen, and also measured changes in enzyme activity through embryonic development (confirmed by Yamashita, 1965). In further studies, the enzyme from fat body was characterized kinetically, and evidence was obtained suggesting activation mechanisms similar to those known from vertebrates. Takehara (1962)

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333

assayed phosphorylase (presumably chiefly from fat body) in overwintering prepupae and in pupae of the slug caterpillar, Monema JEavescens. The pH optimum was about 6.6, the K,,, for glucose-1-P was 11 mM, and activity was greatly increased by the presence of 5’-AMP. The enzyme from lepidopteran fat body (dissected from Samia Cynthia pupae) was further characterized by Stevenson and Wyatt (1964). Activity was maximal near pH 7, and the K,,, values ) for glycogen (about determined for glucose-1-P (about 25 m ~ and 1 g/l) were substantially higher than the corresponding constants for phosphorylase from vertebrate sources. The enzyme also differed from that from vertebrates in being neither activated by cysteine nor inhibited by p-chloromercuribenzoate, so that there was no evidence for essential SH groups. The most interesting observation was the great activating effect of 5’-AMP, and developmental changes in this. Activity measured without AMP was taken to indicate active enzyme and could be expressed as a percentage of total enzyme, or activity found with AMP present. In fat body from diapause pupae, in which glycogen is conserved, active enzyme was only about 1% of the total in Samia Cynthia and as little as 0.2-0.3% in the related Hyalophora cecropia. The fraction of active enzyme was higher in the larva and the adult; it was also elevated in the later stages of development to the adult, which corresponds to the use of glycogen. Increased active phosphorylase was found 1 day after experimental injury to the integument of diapause pupae, also a stimulus to glycogenolysis. Finally, if rinsed fat body dissected from diapause pupae was left in a moist chamber at room temperaturefor 1h, the proportion of activeenzyme rose to 10-30% of the total, while the total level did not change. All these observations suggest a physiologically significant mechanism or mechanisms for interconversion of two forms of phosphorylase, in accordance with the results of Steele (1963) with corpus cardiacum extracts in cockroaches, to be discussed below. Puzzling, however, is the small fraction of total fat body phosphorylase found in the active form in any stage of silkmoth development, which seems to indicate substantial unused enzymic potential. The presence of phosphorylase in insect muscle is implied by Sacktor’s (1955) finding that glycogen supported respiration in housefly flight muscle homogenates as efficiently as free sugars. The enzyme has been demonstrated histochemically in sections of locust muscle by Hess and Pearse (1961), who found that activity was dependent on the presence of AMP and was extremely high in flight muscle’ and much lower in leg muscle.

334

G. R. W Y A T T

In some recent experiments with thoracic muscle of the cockroach, Blaberus discoidalis (S. S . Wyatt and G. R. Wyatt, unpublished), a high level of phosphorylase activity was found. The enzyme was entirely in the soluble fraction of homogenates, and exhibited requirements for glycogen and glucose-1-P similar to those of the enzyme from silkmoth fat body. The cockroach muscle enzyme was activated by AMP, but, in rapidly prepared homogenates, the fraction of total activity that was manifest in the absence of AMP was high-generally between 50% and 90%. During storage of aqueous homogenates, the content of active enzyme declined until after 24 h at 0°C almost none remained, but this change could be inhibited by addition of EDTA and potassium fluoride. Activity assayed with AMP was relatively stable. No precautions that were tried, however, (such as chilling, freezing or anesthetizing the animals with C 0 2 before dissection) significantly altered the high proportion of active phosphorylase found in the initial homogenate. This result seemed unlikely to represent the situation in viuo, since the glycogen (about 0.8% of fresh weight was found) in the muscle would have rapidly broken down, even during rest, and our intent to examine changes in phosphorylase activity with changes in muscular activity was thus frustrated at the outset. Resting frog muscle rapidly frozen yields less than 3% of its phosphorylase in the active a form (Danforth et al., 1962). It is not known whether the high percentage found with the insect was due to exceedingly rapid conversion to the active form during dissection and homogenization, or whether the insect exerts some other type of control over phosphorylase activity in viuo. A few experiments with thoracic muscle from adult cecropia silkmoths gave similar results, although the levels of both phosphorylase and glycogen were much lower, as expected in a moth which burns chiefly lipids in flight. 4. Amylase In contrast to their intracellular metabolic degradation by phosphorylase, the a-1,4 linkages of glycogen and starch can be cleaved hydrolytically by amylase. Amylase has been detected in numerous species of insects as an intestinal enzyme, where its digestive function is obvious. There has also been substantial work on the genetic determination of amylase production in Bombyx mori and Drosophila (Kikkawa, 1953; Kikkawa and Abe, 1960). In silkworms, it is interesting that amylase is found in the hemolymph as well as in the gut, and that the enzymes from these sources are distinct both genetically (Matsumura, 1934, cited by Kikkawa, 1953) and biochemically (It0

BIOCHEMISTRY OF S U G A R S A N D POLYSACCHARIDES

335

et al., 1962). Whereas the intestinal enzyme has maximal activity at pH 9.2 and is somewhat activated by citrate but scarcely affected by halides, the activity found in hemolymph is maximal at pH 64-65, unaffected by citrate, and strongly activated by C1- and Br-. The enzyme in hemolymph is also reported to exhibit a higher ratio of saccharifying activity (release of maltose units) to dextrinizing activity (internal cleavage of the chains) than the intestinal enzyme, but the evidence does not warrant designating them as /?- and a-amylases, respectively. It is also of interest that when an ovary from a strain that has amylase in the blood is implanted into a larva of a strain which normally lacks it, the hemolymph of the host in the late pupal stage is found to contain amylase (Kikkawa, 1950, cited by Kikkawa, 1953). This seems to indicate that the ovary can release amylase or some activator of it into the body fluid. Studies of these amylase mutants has also led to the conclusion that the rise and fall of glycogen in the fat body is unaffected by the amylase level in the hemolymph (Kuroda, 1951, cited by Kikkawa, 1953). Beyond these observations, the source and significance of the hemolymph amylase are unknown. Possibly it may serve to digest glycogen released at times of histolysis, and in this connection it would be interesting to know its changes in activity during the insect’s developmental cycle. In Drosophila melanogaster, amylase activity is controlled by a single genetic locus (Kikkawa and Abe, 1960; Kikkawa, 1960). By the use of zone electrophoresis in agar gel (Kikkawa, 1964), or acrylamide gel (Doane, 1965), stained with iodine after incubation with starch, up to seven amylase zones, or isozymes, have been detected. No more than two appeared in any given homozygous strain. On the basis of substrate specificity and the effects of several inhibitors and activators, all appear to be a-amylases (W. W.Doane, unpublished). The total amylase activity increases progressively from the egg to the late third instar larva, then drops in the pupa and rises again in the adult. In both larvae and adults, blood as well as mid-gut contains high amylase activity, and there are lower levels in extracts of other tissues. The relative activities of the isozymes differ in different tissues. By observing intact intestines embedded in starch-containing gel, Doane (unpublished) has detected the passage of amylase out from the gut lumen through specific sites in the gut wall, and this is presumably the origin of at least some of the enzyme found in the hemolymph. Because of these findings, hydrolysis of glycogen by amylase is indicated in Fig. 2, but it is not at all clear how important or general such a process may be in insect physiology. It should also be noted

336

G . R. W Y A T T

that release of glucose by this pathway would require a further glucosidase to cleave the products of amylase action.

a-

VI. HORMONAL EFFECTSO N CARBOHYDRATE METABOLISM

In insects, as in the vertebrates, the regulation of carbohydrate metabolism is effected in part by hormonal mechanisms, and these have lately been receiving increasing attention. The role of hormones in insect growth and development, including some biochemical aspects, has been discussed in recent reviews (Wigglesworth, 1964; Gilbert, 1964). In Table VI are compiled some experimental observations specifically relating to carbohydrates. The complexity of interactions both among the endocrine glands and in the biochemistry of the receptor systems must be emphasized; thus, when an altered substrate level is observed after extirpation of an endocrine centre, this is but the first step toward establishing an effect of a hormone on an enzyme system. The effects that follow removal of the insect brain or its neurosecretory cells may provisionally be attributed to lack of ecdysone from the prothoracic gland, which is dependent on stimulation by brain hormone; ligation at the thorax could act in the same way. That the conservation of glycogen which follows these operations may be a consequence of the absence of ecdysone is supported by the recent data of Kobayashi and Kimura (1966), who find that 18 h after injection of ecdysone into brainless silkworm pupae (dauerpupae) the incorporation of injected [14C]glucoseinto trehalose is enhanced and its incorporation into glycogen is lowered. A direct effect of ecdysone on glycogen metabolism in the fat body, however, cannot be assumed; it is possible, for example, that the effect on the fat body may be a result of some type of feedback from the enhanced energy demand of stimulated biosynthesis in the epidermis, a tissue known to be sensitive to ecdysone. The consequences of allatectomy are prima facie attributable to lack of juvenile hormone, the only firmly established secretion of the corpora allata (Gilbert, 1964). Again, however, a direct effect on carbohydrate metabolism is doubtful; indeed, Orr (1964), noting especially great accumulation of lipid in the fat body of allatectomized blowflies, suggests that the changes in glycogen may somehow be secondary to altered lipid metabolism. There is other evidence that lipids are especially affected by juvenile hormone (Pfeiffer, 1945), but the primary action or actions of the hormone are unknown, and the impression exists that these may be more closely connected with nucleic

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337

acid and protein synthesis than with catabolic processes. The report of L'HClias (1953, 1955) that sugar is elevated in the tissues and lowered in the hemolymph of allatectomized Dixippus can, unfortunately, not be interpreted since only total reducing substances in heat-dried tissues were determined. By far the most satisfactory evidence for action of an insect hormone on an enzyme system is that obtained with corpus cardiacum extracts and fat body from several species of cockroach. Steele (1961) first reported that injection of aqueous extract of corpora cardiaca into adult cockroaches was followed by elevation of hemolymph trehalose. The activity was impressive, for extract equivalent to as little as 0.002 pair of glands gave 30% elevation of trehalose, larger amounts gave up to 150% elevation, and the increase was demonstrable as early as 30 min after the injection. In further work by Steele (1963) and others (McCarthy and Ralph, 1962; Ralph, 1962; Bowers and Friedman, 1963), it was shown that corpus cardiacum extract can stimulate the release of trehalose when added to isolated fat body in a saline medium, and that the chief source of the released trehalose is fat body glycogen. An enzymic basis for the glycogenolysis was detected in increased activity of glycogen phosphorylase in €at body that had been incubated with cardiacum extract for 2 h, or taken from animals injected with the hormone (Steele, 1963). Hormonally initiated activation of phosphorylase is not surprising in view of the systems known in other animals by which hormones such as epinephrine and glucagon bring about conversion of muscle and liver phosphorylase from inactive to active forms via adenyl cyclase, 3',5'-adenylic acid and phosphorylase kinase w e b s and Fischer, 1962). The hormone from corpus cardiacum is polar, heat-stable and beIieved to be a peptide, and its target organs appear to be fat body and nervous cord, whereas muscle and gut are unaffected. Further analysis of the nature of its effect on phosphorylase has not been reported. In recent experiments, Wiens and Gilbert (1965) have found that when cardiacum extract is added to fat body in oitro, respiration is increased, but at the expense of lipids rather than carbohydrate, for conversion of [14C]glucoseto COz is depressed while oxidation of [14C]acetateand [14C]palmitateis stimulated. They propose that in addition to activating phosphorylase, the hormone may bring about inhibition of phosphofructokinase (thus blocking glycolysis) and activation of trehalose-phosphate synthetase. The increased fatty acid oxidation would then result from compensatory mechanisms related to energy demand. While this is a plausible interpretation of what is

w w W

TABLE VI Hormonal effects on carbohydrate metabolism in insects Endocrine system and experimental treatment

Species and stage*

Brain and prothoracic glands Bombyx mori, P Ligated at thorax Brain removed B. mori, P Ecdysone injected into B. mori, P brainless pupae Neurosecretory cells removed Neurosecretory cells removedt Corpora allata (= CA) CA removed CA removed CA extract injected Fat body inc. with CA extract

Calliphora erythrocephala, A Aedes spp., A Pyrrhocoris apterus, A Phormia regina, A Periplaneta americana, A Blaberus giganteus

Observed effects*

Reference

Use of glycogen is blocked Use of glycogen is blocked Conversion of G to T stimulated, conversion to glycogen inhibited Glycogen accumulates

Ito and Horie, 1957 Kobayashi, 1957 Kobayashi and Kimura,

Glycogen accumulates; lipids low

Van Handel and Lea, 1965

Glycogen accumulates Fat body glycogen slowly rises; so does lipid Blood G falls G uptake increased

Janda and Slama, 1965 Orr, 1964

1966

Thomsen, 1952

c!

e *cl cc

4

McCarthy and Ralph, 1962 Ra ph, 1962

E

0 cl

Corpora cardiaca (CC) CC extract injected

P. americana, A

CC extract injected

P. americana, A

Fat body incubated with CC extract CC extract injected Fat body incubated with CC extract "Injury factor " Integument of diapause pupae injured Sub-oesophageal ganglion Diapause hormone injected

B. giganteus

i 2

T rises, G unchanged, glycogen Steele, 1961, 1963 z falls, phosphorylase activated M McCarthy and Ralph, 1962; T rises, G unchanged Ralph and McCarthy, 1964 p Ralph, 1962 Release of T stimulated

* 0

B. discoidalis Leucophaea maderae, A

T and G rise; glycogen falis Glycogen converted to T; fat oxidized

Bowers and Friedman, 1963 Wiens and Gilbert, 1965

Hyafophora cecropia, P

Blood T raised, fat body glycogen Wyatt, 1961b,1963b;Stevenlowered, phosphorylase acti- son and Wyatt, 1964 vated

B. mori, P

Blood T lowered, ovary glycogen Yamashita and Hasegawa, 1964, 1965; Hasegawa raised and Yamashita, 1965

* *z

Q

U

21

r

2 'p

* Abbreviations as in Table I.

7 The authors point out that the mechanism may not be endocrine, since the same effect is produced by cutting the axons of these Cells.

cl cl

*

z U m vl

w w

\o

340 G. R. W Y A T T seen, other sequences of interaction could be proposed. The system seems readily accessible to further analysis. Superficially similar are certain consequences of integumentary injury in diapausing saturniid silkmoth pupae (Wyatt, 1961b, 1963b). Local wounding of the integument of Hyalophora cecropia or other species in diapause stimulates respiration of the pupa for some days (Harvey and Williams, 1961), but conversion of [l-14C]glucoseto C 0 2 is unaffected (Wyatt, 1963b). Blood trehalose is elevated at the expense of fat body glycogen (Wyatt, 1961b, 1963b), and fat body phosphorylase is converted from an inactive form (i.e. active only with added AMP) to an active form (Stevenson and Wyatt, 1964). These effects cannot depend on the corpus cardiacum, however, for they are observed in pupae from which the brains have been removed without especial care to preserve the allata and cardiaca; indeed, injury has been found to evoke enhanced respiration even in isolated abdomens devoid of known endocrine glands. Therefore, release of an “injury factor” from the site of the wound is postulated, but nothing is known as to its nature. An influence on carbohydrate metabolism in the direction opposite to those just described, namely an accumulation of glycogen at the expense of blood trehalose, has recently been shown in Hasegawa’s laboratory (Yamashita and Hasegawa, 1964, 1965; Hasegawa and Yamashita, 1965). The effect is produced by injecting into silkworm pupae the diapause hormone (which causes production of diapause eggs) prepared from sub-oesophageal ganglia, and the increased glycogen deposition is observed solely in the ovaries. The eggs thus receive the increased reserves needed for overwintering. Nothing is known as to the mechanism of this effect. V I I . GLYCOPROTEINS A N D CHITIN A. GLYCOPROTEINS I N INSECTS

Chitin is a long-chain polymer consisting of ,f3-1,4-linked N-acetylglucosamine together, apparently, with a certain variable percentage of glucosamine. In the insect cuticle, it occurs bound to protein, and the complex is thus appropriately described as a glycoprotein. The extensive studies on its structure by physical and chemical methods have been discussed elsewhere (Richards, 1958; Rudall, 1963; Hackman, 1964). Other glycoproteins in insects have been virtually ignored until very recently. The insect peritrophic membrane contains protein, chitin and

BIOCHEMISTRY OF SUGARS A N D POLYSACCHARIDES

341

other mucopolysaccharide, including, apparently, hyaluronic acid (DeMets and Jeuniaux, 1962; Nishizawa et al., 1963). Mucopolysaccharide was also extracted from mid-guts of Galleria mellonella, purified chromatographically and identified by analysis as hyaluronic acid (Estes and Faust, 1964). Siakotos (1960a, b) separated plasma proteins of Periplaneta americana by paper electrophoresis and, by using stains selective for several prosthetic groups, detected several glycoprotein bands. The carbohydrate content of one of these was sigmficantly elevated during the pre-moult stage. These observations were followed up by the intensive chemical studies of Lipke et al. (1965a). They prepared glycoprotein fractions from P. americana plasma which when hydrolysed released glucose, galactose, mannose, arabinose, xylose, glucosamine and galactosamine. The quantitative proportions of these changed during the moulting cycle, mannose being generally the most abundant component. Fat body contained protein-bound mannose and hexosamine, while extracts of cuticle, upon hydrolysis, released all of the above sugars with the exception of galactosamine. It was established that the pentoses are true polysaccharide constituents and not artefacts of aminohexose degradation. Curiously enough, no sialic or uronic acids, nor fucose, all of which are common in mammalian mucopolysaccharides, could be found. B . METABOLISM OF CHITIN

1. Relations to other carbohydrates In the discussion of glycogen, a relation between this polysaccharide and chitin as reserve nutrients was noted, the quantitative changes in the two being to some extent reciprocal at certain points in the moulting cycle. This was especially brought out by the analyses of Zaluska (1959) on Bornbyx mori throughout its development (Fig. 8). In Hyalophora cecropia, some 8 0 4 5 % of the integumentary chitin is digested and resorbed during the pupal moult (Bade and Wyatt, 1962; Fig. 9), and again during the adult moult (Passoneau and Williams, 1953). The further role of chitin as a reserve nutrient apart from the moulting process is emphasized by Locke (1964), who notes that in Rhodnius and some other insects resorption of the endocuticle occurs during prolonged starvation, and deposition of endocuticle follows feeding. The metabolic mobility of the cuticle is supported by the evidence obtained by Lipke et al. (1965b) that in Periplaneta americana the synthesis and degradation of cuticle polysaccharide is continuous throughout the moulting cycle.

342 G . R. W Y A T T Some evidence concerning precursors and reserves for chitin synthesis has been obtained with radioisotopes. Bade and Wyatt (1962) injected 14C-glucose into cecropia silkworms at the end of the fourth instar and showed that by the end of the feeding period in the fifth instar the retained radioactivity was very largely in the cuticle. The

Time from spinning (days)

Fro. 9. Content of carbohydratesand lipids in the cecropia silkworm from the mature larva to the pupa in diapause. 0 ,Sugars; 0, chitin; A,glycogen; 0 , total of estimated carbohydrates (sugars chitin glycogen); A,lipids. Day 0 represents the beginning of cocoon spinning; S, period of spinning; P, pupal moult. (From Bade and Wyatt, 1962.)

+

+

subsequent new pupal cuticle, taken 2 days after pupation, was again highly radioactive, the specific activity of its chitin being close to that of the chitin of mature larvae and much greater than that of their glycogen, free sugars or lipids. This appears to indicate specific transfer of material from the larval to the pupal cuticle. Such transfer might occur by products of digestion of the old cuticle being directly incorporated into the new as they are resorbed. The possibility of specific

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internal stores, perhaps in glycoproteins, was also considered, since much of the pupal endocuticle is deposited after the moult. The possible role of hemolymph and tissue glycoproteins as chitin precursors was tested in the cockroach, P . americana, by Lipke et al. (1965b). When [14C]glucosewas injected before a moult, the greatest incorporation was into blood trehalose and cuticle polysaccharides (including chitin). Mucopolysaccharides of plasma and fat body acquired relatively little radioactivity, and no evidence was obtained that any such fraction was a significant precursor for cuticle synthesis. Cuticular hexosamine appeared to be formed endogenously, and blood trehalose qualified kinetically as its chief precursor. The resolution of the discrepant implications of these two sets of experiments is not clear, but perhaps there are real differences between the two types of moult; the cockroach was growing and would require new synthesis to make a new integument larger than the old, whereas during silkworm pupation the pupal cuticle even when fully formed contains only about half as much chitin as the mature larval cuticle, and its deposition entirely at the expense of the latter would be possible.

2. Synthesis of chitin Despite the dominant place of chitin in the arthropod economy, very little has been established concerning the enzymes that make it. The synthesis of chitin from UDP-acetylglucosamineby a cell-free preparation from the mould, Neurospora crassa, was achieved by Glaser and Brown (1957). Activity was located exclusively in a particulate fraction recovered from homogenates by centrifugation at 140000 x g, and incorporation from UDP-[14C]acetylgluc~~amine into insoluble chitin was demonstrated. Soluble chitodextrins were required as a primer, and activity was strongly stimulated by the presence of N-acetylglucosamine. Up to 20% of the precursor radioactivity was incorporated during 2.5 h incubation, and net synthesis was also shown by chemical measurement of insoluble hexosamine. The enzymes of chitin synthesis in locusts were sought by Candy and Kilby (1962). In Schistocerca greguria, they demonstrated by direct analysis and by in uiuo isotope incorporation that the legs and wings are active in chitin synthesis for approximately 7 days after the adult moult. Wings from 2-3-day-old adults were therefore used as an enzyme source, and all of the enzymic activities required to convert glucose to UDP-acetylglucosamine were demonstrated. Attempts to detect activity of chitin synthetase itself, however, which would incorporate the latter compound into chitin, were unsuccessful, although the 13-kA.I.P. 4

344

G . R. W Y A T T

extracts were fortified and conditions were varied in a number of ways. Evidence for chitin synthetase in the southern armyworm, Prodeniu eridania, has more recently been obtained by Jaworski et ul. (1963) and Porter and Jaworski (1965). Radioactivity was incorporated from UDP-[14C]acetylgluc~~amine into insoluble material, from which it could be partially released by the action of chitinase. In addition to the high-speed particulate fraction, the mitochondria1 fraction apparently contained activity (so did the debris residue, but this could be due to unbroken cells). Activity was greatest in late final instar larvae and prepupae, as expected; it appears, however, that the best incorporation was about 1% of the labelled precursor during 2 h incubation. Acetylglucosamine was included in the incubations, but the effect of its omission is not stated. Similar results with crustacean enzymes are reported by Carey (1965) : a high-speed particulate preparation from Artemia salina or epidermis of blue crabs incorporated about 1% (in one instance 4%) of radioactivity into a product identified as chitin. It is of incidental interest to note that the content of UDP-sugar derivatives in the hemolymph and tissues of cecropia silkmoth pupae has been examined (Carey and Wyatt, 1960). UDP-glucose and UDPacetylglucosamine were found, but, surprisingly, the most abundant compound in the hemolymph was identified as UDP-acetylgalactosamine. This is presumably a precursor of mucopolysaccharides. In wing epidermis, the content of these nucleotides rises sharply at the beginning of adult development, but the proportions of the individual compounds at this time were not determined. 3. Enzymic degradation of chitin The digestion of the chitin (along with the proteins) of the endocuticle is a central feature of the arthropod moult. The process has not been extensively studied, and what has been found can be summarized only briefly here. The studies of Jeuniaux have been presented in a monograph (Jeuniaux, 1963). Evidence is presented for two steps in the digestion of chitin : chitinase

chitobiase

Chitin-chitobiose-N-acet (+chitotriose acetylglucosamine)

y lglucosamine

+

Chitinase is assayed by loss of turbidity in a suspension of chitin, and chitobiase by colorimetric estimation of acetylglucosamine. Chitinase is produced by Streptomyces antibioticus grown on colloidal chitin as

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carbon source, and has been purified by a procedure of which a key step is adsorption onto the insoluble substrate, chitin, and release by digestion of the latter. A chitinase free of chitobiase has also been obtained by fractionation of cockroach gut extracts (Powning and Irzykiewicz, 1963). Chitobiase can be demonstrated in various sources, such as almond emulsion and silkworm hemolymph, as well as Streptomyces culture fluid. Some studies have been carried out on distribution of chitinolytic activity in insect tissues, and changes during the moulting cycle. In Bombyx mori, “total chitinolytic activity” (assayed by production of acetylglucosamine from finely divided chitin) in extracts of epidermis is very low in the intermoult, and rises to a sharp peak during larval and pupal moults (Jeuniaux, 1961). In the pupa, both chitinase and chitobiase appear with the moulting fluid in the second half of the pupal stage, and disappear with resorption of this fluid just before emergence of the adult (Jeuniaux, 1963). This resembles the previous findings of Passoneau and Williams (1953) for Hyalophora cecropia, except that in cecropia, during the first two-thirds of adult development, moulting fluid was present as a gel which conteined little activity and then liquefied and became enzymatically active. Both workers, and also Zielinska and Laskowska (1958), found substantial levels of amino sugars in the moulting fluid. Waterhouse et al., (1961) have investigated chitinase activity in termites and the cockroach, Periplaneta americana. The assay conditions were studied, and it was shown with extracts from several cockroach tissues that the products from chitin were N-acetylglucosamine plus small amounts of glucosamine. Activity in deacetylating acetylglucosamine to glucosamine, however, could not be demonstrated. Among the tissues of the adult cockroach, the highest chitinase activity was found in the integument, but substantial activity was also present in the gut, hemolymph and saliva (Waterhouse and McKellar, 1961). In roaches as much as 200 days old, chitinase was still highly active, though lower than in young adults. The function of such widespread chitinase is not obvious; one possibility is that it may act in the resorption of chitin during starvation, as mentioned above.

VIII. GLYCEROL A N D SORBITOL A specialized conversion that occurs in the overwintering stage of a number of insects is that of glycogen to polyhydric alcohols, which can be subsequently largely reconverted to glycogen. This was dis-

346 G . R . WYATT covered independently in the diapause embryo of Bombyx mori, which produces sorbitol and glycerol (Chino, 1957b, 1958), and in the cecropia silkmoth pupa, which produces glycerol (Wyatt and Meyer, 1959; see also Fig. 1). Although glycerol production in these species coincides with the onset of diapause, glycerol can be produced by non-diapausing insects, as illustrated by its appearance in adult carpenter ants upon exposure to cold (Dubach et al., 1959; Tanno, 1962). Glycerol production could also account for disappearance and reappearance of glycogen in overwintering adults of the lady-bird beetle, Semiadalia undecimnotata (Hodek and Cerkasov, 1963). The high level of 5 molal (about 25% of body weight) has been recorded for diapause prepupae of the wheat-stem sawfly parasite, Bracon cephi (Salt, 1959). The production of glycerol under various conditions has been carefully studied in the slug caterpillar, Monemaflauescens (Takehara and Asahina, 1961;Takehara, 1963a, b, 1964). Ecologically, the accumulation of glycerol and sorbitol appears to be connected with resistance to cold, and they probably act by lowering the supercooling point rather than by conferring the ability to survive freezing (Salt, 1961; Ssmme, 1964, 1965a, b; Asahina, 1965). The biochemical pathway of glycerol production is presumably via reduction from the triose level of glycolysis, and appropriate enzymes have been demonstrated in insects (Faulkner, 1956, 1958; Chino, 1960, 1961), but the precise sequence of reduction and dephosphorylation has not been established. Most likely is reduction of dihydroxyacetone phosphate to a-glycerophosphate, for glycerophosphate dehydrogenase is highly active in insect tissues. Metabolic control of glycerol production at this step, which was at first postulated because of the degeneration of the cytochrome system during diapause (Wyatt and Meyer, 1959),now seems unlikely for reasons that have been discussed (Harvey, 1962; Wyatt, 1963a), and it is suggested that the dephosphorylation to free glycerol may be physiologically regulated. This would imply a specific a-glycerophosphatase, and preliminary evidence for such an enzyme in the pupal cecropia fat body has been obtained (B. P. Thomas and G. R. Wyatt, unpublished experiments). It is of interest in this connection that exposure of wax moth larvae (Galleria mellonella) to cold results in lowered a-glycerophosphate and elevated inorganic phosphate (Lenartowicz, 1961). ACKNOWLEDGEMENTS

I am indebted to Drs. R. F. Derr and M. Kobayashi for allowing me to see their manuscripts before publication, and to Dr. Winifred W.

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Doane for permission to cite unpublished results. Dr. Doane also assisted by reading the manuscript. Work in my laboratory, and the preparation of this paper, were supported by the Whitehall Foundation and the National Institutes of Health, U.S. Public Health Service (grant no. AI-01028).

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Evans, W. A. L. and Marsden, J. (1956). Carbohydrases in blowfly larvae. Nature, Lond. 177, 478. Evans, W. A. L. and Payne, D. W. (1964). Carbohydrases of the alimentary tract of the desert locust, Schistocerca gregaria. J. Insect Physiol. 10, 657-674. Fairbairn, D. (1958). Trehalose and glucose in helminths and other invertebrates. Can. J. Zool. 36, 787-795. Fairbairn, D. and Passey, R. (1957). Occurrence and distribution of trehalose and glycogen in the eggs and tissues of Ascaris lumbricoides. Expl Parasit. 6, 566-574. Faulkner, P. (1 955). A hexose-1-phosphatase in silkworm blood. Biochem. J. 60, 590-596. Faulkner, P. (1956). Enzymic reduction of sugar phosphates in insect blood. Biochem. J. 64.436-441. Faulkner, P. (1958). Polyol dehydrogenase of the silkworm. Biochem. J. 68, 374-380. Florkin, M. (1937). Variations de la composition du plasma sanguin au cours de la mttamorphose du ver & soie. Archs int. Physiol. 45, 17-31. Florkin, M. and Jeuniaux, Ch. (1964). Hemolymph: composition. In “The Physiology of Insecta” (M. Rockstein, ed.), Vol. 3, Chap. 2. Academic Press, New York. Florkin, M. and Jeuniaux, Ch. (1965). MCtabolisme du trChalose et du glycogkne chez le ver B soie, en relation avec la mue, le filage et les mttamorphoses. Bull. Acad. r. Belg. Cl. Sci. 51, 541-552. Frerejacque, M. (1941). Trthalose et trChalase. C. r. S6anc. SOC.Biol. 213, 88-90. Friedman, S. (1960a). The purification and properties of trehalase isolated from Phormia regina Meig. Archs Biochem. Biophys. 87,252-258. Friedman, S . (1 960b). Occurrence of trehalose-6-phosphatase in Phormia regina Meig. Archs Biochem. Biophys. 88, 339-343. Friedman, S. (1961). Inhibition of trehalase activity in the hemolymph of Phormia regina. Archs Biochem. Biophys. 93, 550-554. Geigy, R., Huber, M., Weinman, D. and Wyatt, G. R. (1959). Demonstration of trehalose in the vector of African sleeping sickness: the tsetse fly. Acta trop. 16, 255-262. Gelperin, A. (1966). Control of crop emptying in the blowfly. J. Insect Physiol. 12, 331-345. George, J. C. and Nair, K. S. S. (1964). Quantitative study of changes in fat and glycogen during metamorphosis of the dermestid beetle, Anthrenus vorax. J. Anim. Morph. Physiol. 11, 162-172. Gilbert, L. I. (1964). Physiology of growth and development: endocrine aspects. In “The Physiology of Insecta” (M. Rockstein, ed.), Vol. 1, chap. 5. Academic Press, New York. Gilby, A. R., Wyatt, S. S. and Wyatt, G. R. (1966). Trehalases from the cockroach, Blaberus discoidalis: activation, solubilization and properties of the muscle enzyme and some properties of the intestinal enzyme. Acra biochim. polo. (in press). Gilmour, D. (1961). “The Biochemistry of Insects”. Academic Press, New York. Glaser, L. and Brown, D. H. (1957). The synthesis of chitin in cell-free extracts of Neurospora crassa. J. biol. Chein. 228, 729-742. Grace, T. D. C. and Brzostowski, H. W. (1966). Analysis of the amino acids and sugars in an insect cell culture medium during cell growth. J. Insect Physiol. 12, 625-633. 13.

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Author Index Numbers in italics are the pages on which the references are listed

A Abderhalden, F., 148, 187 Abe, K., 334,335,354 Acree, F., 155, 187 Adkisson, P. L., 146, 190 Adrian, R. H., 3, 27 Agarwal, H. C., 143, 168, 171,187 Aidley, D. J., 24, 25, 27, 27 Albrecht, G., 89, 187 Allais, J. P., 117, 144. 187 Allen, R. R., 209,210 Allmann, L. K., 311,347 Altman, P. L., 291, 347 Alvarado, F., 303, 358 Amin, E. S.,93, 187 Anderson, A. D., 37, 39, 40, 61 Anders, G., 92, 187 Andersen, S. O., 216, 234,271, 272, 280 Anderson, T. F., 222,265,280,285 Ansell, G. B., 135, 138,188 &on, W. J., 301, 355 Aruga, H., 56, 67 Asahina, E., 296, 346, 347, 358 Asakura, S., 214,280 Ashworth, C. T., 98,188 Asperen, K. van, 111,188 Auclair, J. L., 43, 49, 61, 64 Avigad, G., 315, 347 Ax, H., 183, 200

B Babcock, K. L., 117, 188 Babcrs, F. H., 326, 347 Bachmann-Diem, C., 43,62 Bade, M. L., 131, 150, 175, 188, 327, 328, 329, 341, 342,347 Baker, F. C., 39, 53, 61,66 Baker, F. D., 98, 188 Baker, G., 153, 188 Baker, G. L., 153, 155,188 Baldwin, E., 40,61

Ball, E. G., 176, 188 Ball, H. S., 257, 280 Baranska, J., 209, 211 Barbier, M., 92, 93, 168, 186, 188, 193, 202

Barker, R. J., 164,195 Barlow, J. S.,93, 95, 188 Barrington, E. J. W., 288, 347 Barron, E. S. G., 110,188 Bartel, A. H., 44,61, 64 Bartos, Z., 326,354 Bataillon, E., 327, 347 Bayer, Sh., 125,200 Bayley, S. T., 266, 285 Beall, G., 86, 188 Beament, J. W. L., 152, 153, 188, 277, 280 Becht, G., 14, 17, 27, 27 Beck, S. D., 146,168,173,175,187,190, 191,204 Beckmann, R., 186,190 Beenakkers, A. M., 125, 126,188, I89 Beer, M., 263, 281 Belton, P., 4, 27 Ben-Hayyim, G., 215,263,279,280 Bennett-Clark, H. C., 260, 268, 277, 280 Bennett, A. F., 209, 211 Berger, J. E., 184, 206 Bergerard, J., 117, 144, I87 Bergmann, E. D., 159, 199, 209,210 BergmaM, W., 93, 168, 189 Bernard, C., 325,347 Beroza, M., 186,197 Berridge, M. J., 48, 58, 61 Berthelot, M., 290,347 Bettini, S., 123, 189 Betts, E., 209, 210 Betz, N. L., 327,355, Beutler, R., 295, 347 Bhakthan, N. M. G., 111, 116,189,194

362

AUTHOR INDEX

Bhat, J. V., 131, 142, 167, 174,204,205, 298, 306, 357 Bheemeswar, B., 43,6I Biale, J. B., 138, 189 Bieber, L. L., 143, 189 Biedermann, W., 221,280 Birch, G. G., 290, 347 Bjerke, J. S., 183, 205 Blackith, R. E., 146, 189 Blewett, M., 145, 159, 193, 194 Bloch, K., 129, 130, 131, 157, 159, 161, 167, 172,189,191,192,194 Bloomfield, D. K., 129, 131, 189 Blum, M. S., 89, 92, 189,198 Boccad, M., 123,189 Boch, R., 169, 189 Bode, C., 119,189 Bodenstein, D., 47, 62 Bodenstein, D., 185, 189 Boell, E., 117, 189 Boettiger, E. G., 6, 14, 18, 29, 30, 124, 189 Boettiget, E. G., 18, 20, 29 Bogdanovsky, D., 92, 93, 168, 186, 202 Bogdanowsky, D., 168,188 Boistel, J., 20, 28 Bonner, J., 266, 280 Booth, F., 214, 278, 284 223, 225, 228, 280 Bouligand, Y., Bowers, W. S., 154, 180, 190, 210, 210, 337,339,347 Boyle, P. J., 2, 28 Brazel, J. R., 90, 190 Bressler, R., 126, 190 Bricteux-Grkgoire, S.,299, 305,347,348 Bridges, R. G., 143, 144, 190,192 Brierley, G., 137, 143, 144, 193 Brighenti, A., 40, 62 Brookes, V. J., 143, 189, 209, 210 Brown, A. .W. A., 37, 38,41,42,43,44, 46, 53, 54, 62, 143, 193 Brown, D. H., 343,351 Brown, G. W., 42, 62 Brown, W. H., 92,190 Bruner, J., 262, 280 Brunet, P., 276, 280 Bruno, C. F., 35, 37,62 Bryce, B. M., 167,175,197 Brzotowski, H. W., 302, 303, 351

Buchanan, J. M., 40,62 Bucher, N. L. R., 176,202 Bucher, T., 297, 318,348 Bucher, Th., 119, 190, 198, 208 Buck, J. B., 41,62 Bulbring, E., 23, 28 Bull, D. L., 146, 190 Burden, G. S., 155, 187 Burge, R. E., 214, 278,284 Burks, M. L., 209,210 Bursell, E., 44, 45, 52, 53, 62 Butenandt, A., 92, 169, 186, 190 Butler, C. G., 186, 190

C Cabib, E., 307, 348 Cadenas, E., 303, 358 Callow, R. K., 186, 190 Candy, D. J., 263,280,306,307,343,348 Carey, F. G., 141, 142, 190, 208, 263, 280, 301, 344, 348,360 Carlisle, D. B., 161, 193, 209, 210 Carlstrom, D., 216, 217, 220, 280 Carpenter, W. D., 150, 190, 329,348 Casida, J. E., 168, 171, 175, 187, 190 Castle, E. S., 263, 266, 280 Ceccarini, C., 315, 348 Cerf, J. A., 9, 10, 20, 21, 22, 28 Cerkasov, J., 346, 352 (I Chad, I., 215, 263, 279, 280 Chadha, M. S., 169,191 Chadwick, L. E., 124,191 Chajkinowna, S., 37, 67 Chamberlain, J. C., 89, 148, 194 Chambers, D. L., 153,199 Chance, B., 119,191 Chang, C. K.,1293, 312, 320, 323, 327, 348 Changeux, J. P., 309,355 Chauvin, R., 33, 46, 62, 159, 191 Chararas, C., 311, 320, 348,349 Chefurka, W., 43, 65, 70, 191, 288, 302, 348 Cheldelin, V. H., 125, 143, 189, 196 Chen, P. S., 43, 62 Chibnall, A. C., 93, 191 Chino, H., 89,90,99,100,101,102,103, 104,105, 106, 107, 108,109,112,113,

363

AUTHOR INDEX

114,115,139, 140,148,149,151,191, 195, 327, 346, 348,349 Chippendale, G. M., 146, 191 Chojnacki, T., 142, 191 Clark, A. J., 130, 159,161, 172,189,191, 192 Clark, G. L., 220, 266,280 Clayton, R. B., 70, 157, 161, 163, 167, 171, 172, 175, 176, 178, 192, 209,210, 210,211 Clegg, J. S.,297,303, 304, 306,317,321, 324, 325,349 Cleland, M., 176, 208 Clement, G., 175,192 Clements, A. N., 148,192,302,306,330, 348 Clyton, R. B., 175, 188 Cochran, D., 35, 62 Cochran, D. C., 37,62 Cohen, C. F., 164, 195 Cohen, P. P.,42, 62 Cole, M. J., 175, 190 Colla, A., 40,62 Colvin, J. R., 215, 263, 281 Condoulis, W. V., 251, 263, 264, 265, 268,281,348 Constantin, M. J., 113, 192 Conway, E. J., 2, 28 Cordero, S. M., 37, 38,62 Cori, C. F., 334,349 Cornforth, J. W., 157, 192 Cornforth, R. H., 176,202 Conerbe, J., 42, 63 Cottrell, C. B., 267, 268, 281 Coupland, R. E., 141,192 Courtois, J. E., 311, 314, 315, 320, 322, 348,349 Couvreur, E., 327,347, 349 Cowan, S., 12,29 Cox, J. T., 143, 190 Cox, M. E., 97, 153, 154, 195 Craig, R., 34, 40, 44,61, 62, 64 Crane, F. L., 166, 199 Cromartie, R. I. T., 157,192 Crone, H. D., 144,192,209,210 Cross, A. D., 210,211 Crowe, P. A. Sr., 327, 328,355 Currey, J. D., 220,281 Cmrnovsky, C. von, 294, 295,349

D Dadd, R. H., 145, 146, 160, 192, 303, 349 Dahlquist, A., 315, 322, 349 Daly, H. V., 267, 281 Danforth, W. H., 330, 334,349 Daniel, L. J., 135, 140, 207 Danon, M. D., 214,284 Dateo, G. P.,47, 66 Davis, G. R. F., 42, 62 Day, M. F., 222,283,297, 320,359 Debris, M. M., 311, 320, 348, 349 Debye, P.,274, 281 de Harven, E., 17,28 Delaunay, H., 51, 62 Delbruck, A., 119,208 del Castillo, J., 9, 10, 13, 14, 20, 28 DeLuca, H. F., 137, I94 DeMets, R., 341, 350 Demyanovsky, S. Ya., 86, 87, 192 Dennel, R., 227,264,281 Derr,R. F., 291,292,296,298,310,311, 313, 314,322,350,356 Desai, R. M., 37,40, 42, 43, 62, 63 Desnuelle, P.,98, 113, 115, 116, 192 Dethier, V. G., 290, 291, 294, 299, 350 Deuel, H. J., 71, 193, 328, 350 De Waart, J., 43, 67 Dick, A., 119, 203 DiNella, R. R., 113, 193 Ditman, L. P., 116, 193 Dittmer, D. S., 291, 347 Doane, W. W., 335,350 Dole, V. P., 103, 193 Domroese, K. A., 84, 87, 88, 99, 100, 101, 112, 113, 114, 115, 120, 122, 123, 124, 193,195, 329,350 Doudoroff, M., 310,350 Downing, D. T., 193 Drach, P., 225, 227, 281 Dresden, D., 27,27 Dubach, P., 346,350 DuchPteau, G., 13, 28, 37, 38, 63 Duchateau, Gh., 293, 294, 295,350 Duchgteau-Bosson, G., 293, 312, 323, 350 Dudel, J., 14, 28 Dumm, M. E., 138,202 Duperon, P., 168,193

364

AUTHOR I N D E X

Duspiva, F., 98, 193, 311, 319, 347, 350 Dutky, R. C., 161, 171, 175, 193, 197, 202 Dutrieu, J., 296,299, 324,350

E Eapen, J., 111, I94 Earle, N. E., 209,210 Edman, K. A. P., 24,28 Edwards, A.M., 167,171,175,192,210, 211 Edwards, G. A., 17,28 Eggerer, H., 166, 200 Egorova, T. A., 293,296, 306,350 Eguchi, G., 214,280 Ehrhardt, P., 291,292,311,321,322,350 Eisner, T., 98,99, 169,191,193,209,211 Elliott, G. F., 246, 271, 281 Ellis, P. E., 161, 193, 209, 210 Emmerich, H., 43, 63 Engbaek, L., 13, 14,28 Estes, Z. E., 341, 350 Etienne, J., 117, 144, 187 Evans, D. R., 290, 291, 294, 297, 299, 303,304, 306, 317, 321,349,350 Evans, W. A. L., 311,322,350 Ewer, D. W., 17, 30

F Fairbairn, D., 291, 324, 329,351 Falk, M., 214,283 Farber, L., 43, 62 Farkas, T., 96, 196 Fast, P., 209, 210 Fast, P. G., 70, 81, 89, 91, 143, I93 Fatt, P., 5 , 9, 15, 20, 28 Faulkner, P., 301, 306, 346, 351 Faust, R. M., 341,350 Fawzi, M., 92,96,193 Feist, E., 147, 193 Felauer, E., 92, I90 Feller, D. D., 147, I93 Feng, H., 293, 311, 312, 320, 323, 324, 327,348,355 Filosa, M. F., 324, 325, 349 Fischer, E. H., 337, 354 Fleischer, S., 137,138,143,144,193,195 Florkin, M., 13, 28, 33, 37, 38, 63, 99,

193,291, 293,294,295, 299, 305, 312. 323,347,348,350,351 Fodor, P. J., 115, I93 Folkers, K., 137, 145, 166, 206 Folley, S. J., 147, 202 Forchielli, E., 276, 282 Forgash, A. J., 40, 65 Fowler, K. S., 139, 199 Fox, A. S., 37, 65 Fraenkel, G., 49, 63, 130, 145, 159, 160, 189, 193, 194, 202,206,217,218,219, 266,267,269, 271,273,281, 295,351 Frank, G. B., 24,26,28 Frappez, G., 37, 63 Fredrickson, D. S., 102, 103, 116,194 Frei, E., 217, 281 Frerejacque, M., 310, 351 Freure, E., 92, 190 Freure, R. J., 92, 190 Frey-Wyssling, A., 214,278,281 Friberg, U., 227,286 Fried, J. H., 210,211 Friedberg, S. J., 126 Friedman, S., 301, 307, 310, 313, 314. 322, 324, 337, 339,351 Friend, W. G., 145,194 Frisch, A. M., 175,192 Fulton, R. A., 89, 119, 148,194 Furlenmeier, A., 210, 210 Fiirst, A., 210,210 Fuseau-Braesch, S., 157, 194

G Gaeta, I., 98, I94 Garattini, S., 157, 194 Garbus, J., 137, 194 Garcia, I., 42, 63 Gebhardt, W., 221,281 Geigy, R., 296,351,360 Gelperin, A., 298, 351 George, J. C., 111, 116, 140, 150, 194, 201, 327, 328,351 Gibson, D. M., 132,202 Gilbert, L. I., 70, 81, 83, 84, 86, 87, 89, 90,93,99,100,101,102,103,104,105,

106, 107, 108, 109, 112, 113, 114, 115, 119, 120,122, 123,124,140,141,142, 146, 148, 149, 151, 157, 161, 167, 168, 174, 175, 176, 178, 179, 180, 181, 182,

AUTHOR INDEX 183,184,186,191,193,194,195,200, 204, 329, 336, 337, 339, 350,351, 359 Gilby, A. R., 70, 97, 153, 154, 155, 195, 207, 310, 313, 314, 315, 316, 319,351 Giles, C. H., 275, 281 Gilmour, D., 42, 63, 70, 124, 191, 195, 263,281,299,351 Glascock, R. F., 147,202 Glaser, L., 343,351 Glass, E. H., 145, 203 Glassman, E., 37, 64 Glimcher, M. J., 277,281 Gloor, U., 166, 195 Goldberg, M., 275, 276,282 Goldsmith, T. H., 164,195 Goodfellow, R. D., 161, 167, 168, 174, 175, 176, 178, 179, 180, 181, 182,194, 195 Goodman, R. M., 301,359 Goodwin, T. W., 157,164,195,256,282 Gordon, H. T., 145,195 Gordon, R. S. Jr., 102, 103, 116,194 Grace, T. D. C., 302, 303,351 Grainger, M. M., 263,275,283,341,354 Graves, B., 341, 343,354 Gray, H. E., 49,63,295,351 Green, D. E., 137, 138,195 Grundfest, H., 4, 5, 6, 8, 9, 10, 12, 18, 20, 21, 22, 27, 28, 30,31 Gudalina, N. G., 327, 358 Guibourt, 290, 352 Gupta, B. L., 118,201 Gupta, P. D., 47, 51, 63, 67 Gussin, A. E., 140, 195 Gussin,A. E. S.,313,314,315,316,319, 329,352

H Haahti, E., 73, 195 Habibulla, M., 140, 141, 142, 195 Hack, M. H., 140,195 Hackman, R. H., 271, 273, 275, 276, 282,286,288, 340, 345,352,359 Hadorn, R., 37,39, 67 Hagiwara, S., 3, 28 Hamamura, Y.,160,195 Hamlin, J. T., 103, 193 Hanbury, D., 290,352 Handa, N., 341,355

365

Hanes, C. S., 332,352 Hansen, K., 314, 315,352 Hansen, O., 290, 292, 295, 352 Happ, G. M., 209,211 Harker, J. E., 244,282 Harlow, P. A., 12,28 Harrington, J. S.,43,48,49, 63 Harvey, W. R., 340,346,352 Harwood, R. F., 96,195 Hasegawa, K., 339, 340,352, 360 Hassan, A. S. A., 275,281 Hassemer, M. M., 327, 328, 355 Hassid, S. Z., 310, 350 Hastings, E., 153, 188 Hastings, J. W., 239, 285 Haub, J. G., 53, 64, 150, 196 Hawthorne, J. N., 135, 138,188 Hayachiya, K., 160,195 Hayashi, Y., 37,41,63 Haydack, M. H., 47,63 Hecker, E., 186,190 Hedin, P. A., 43, 51, 65 Heed, W. B., 169,196 Heller, J., 40,63,167,196,289,293,299, 352 Helmreich, E., 334, 349 Hemming, U., 166, 200 Hemmingsen, A. M., 289,352 Henderson, W. F., 218,219,286 Hennebeny, G. O., 147,200 Henry, S. M., 147,196 Herb, U., 210,210 Herodek, S., 96, 196 Herz, R., 25, 31 Herzog, R. O., 219, 282 Heslop, J. P., 138, 196 Hess, R., 330, 333, 352 Hestrin, S.,214, 284 Hill, D. L., 117, 196 Hill, E. P., 315, 324, 352 Hill, R. B., 9, 11, 12, 28 Hines, W. J. H., 148, 196, 306, 352 Hinton, T., 42,63 Hirano, C., 161, I97 Hitchcock, F. A., 53, 64, 150, 196 Hobson, R. P., 157, 196 Hocks, P., 179,198,210,210 Hodek, I., 346, 352 Hodgkin, A. L., 2, 3,4,5,21,24,28,29

366

AUTHOR INDEX

Hodgson, E., 142,196 Hoffmeister, H., 179, 197, 198 Hofmann, F., 148,196 Hokin, L. E., 135, 138, I96 Hokin, M. R., 135, 138, 196 Hollmann, S.,303, 352 Holman, M., 23,28 Hopf, H. S., I96 Hoppe, W., 179,197 Horie, Y.,299, 302, 314, 320, 332, 338, 352,353 Homing, F. C., 171,206 Homing, M., 176,202 Homing, M. G., 73,196 Horowicz, P., 2, 3, 5, 24, 29 Hoskins, D. D., 125, 196 Hoskins, W. M., 40,64 House, H. L., 97, 145, 196, 288, 303, 353 Houwink, R., 266,282 Houx, N. W. H., 143,187 Howden, G. F., 146,189,289,290,292, 311, 322,353 Hoyle, G., 3,6,7,8,9, 10, 11, 13, 14, 17, 20, 21, 22, 24, 26, 27,27, 28, 29 Hsu, S., 118, 197 Huber, L., 214,283 Huber, M., 296,351 Huber, R., 179,197 Hiibscher, G., 134,197 Huckel, E., 274,281 Hudson, B. W., 44,61, 64 Hiigel, M. F., 168,193 Hulsmann, W. C., 128, I97 Hummel, H., 179, 198 Humphrey, G. F., 292,353 Huxley, A. F., 21,29,246,271,281 Huxley, H. E., 214,282

I Iino, T., 214,280 Ikeda, K., 17,20,29 Ikekawa, N., 174,201,203 Imms, A. D., 45,64 Irreverre, F., 52, 53, 64, 67 Irzykiewicz, H., 37, 39, 41, 64 Ishii, S., 159, 161, 197, 311, 360 Isselbacher, K. J., 98, 135, 197 Iterson, G. van, 266, 282

Ito, T., 41, 56, 64, 160, 197, 302, 303, 332, 335, 338,353

J Jackson, S. F., 214, 278,284 Jacob, F., 309, 355 Jacob, M. I., 132,202 Jacobson, M., 180, 186, 197, 208, 210, 210 Jakus, M. A., 223,282 Janda, V. V. Jr., 338,353 Jaworski, E., 263,282, 344,353 Jaworski, E. G., 150,190,209,211,329, 344,348,356 Jeanloz, R., 276, 282 Jenden, D. J., 6, 29 Jenny, N. A., 173,204 Jensen, M., 215, 218, 219, 243,282 Jeuniaux, Ch., 99, 193, 288, 291, 293, 299,305,312,323,341,344,345,347, 348,350, 351,353 Jewell, B. R., 25, 29 Jezewska, M. M., 40, 63 Joel, C. D., 176,188 Johannson, B., 148,197 Johnson, L. H., 153,188 Johnston, J. M., 98, 188 Johnston, N. C., 92, 168, 186,190,207 Jones, W. A., 186,197

K Kalf, G. F., 140,208,290,293,294,301, 310, 313, 314, 353,360 Kallen, R. G., 132, 197 Kamienski, F. X., 209,210 Kandler, O., 291, 353 Kaplan, N., 310,350 Kaplanis, J. M., 171, 206 Kaplanis, J. N., 131, 161, 167, 171, 172, 175, 179, 185, 193, 197,199,202,203, 206 Karlson, P., 179, 197, 198 Katz, B., 4, 5 , 9, 13, 15, 28, 29 Katzen, H. M., 326, 358 Kawahara, F. S., 170, 205 Kawashima, K., 160, 197 Keith, A. D., 209,210 Kellenberger, E., 214, 283 Keller, E. C., 37, 64

AUTHOR INDEX

Kelly, F. C., 214, 278, 284 Kennedy, E. P., 124, 134, 135,198, 207 Kent, P. W., 269,282 Kerkut, G. A., 6, 12,29,30 Kerly, M., 302, 353 Kerur, D., 145,206 Kessell, I., 166, 200 Keynes, R. D., 5, 30 Kikkawa, H., 334, 335,353,354 Kilby, B. A., 37, 40, 42, 43, 62, 63, 64, 70, 89, 198, 263, 280, 288, 289, 290, 292, 306, 307, 311, 322, 330, 343, 348, 353,354 Kimura, S., 336, 338,354 Kinsella, J. E., 209, 210,211 Kirirnura, J., 177, 198 Kirscher, H. W., 169, 196 Kiyasu, J. Y.,135,207 Kleiber, M., 147, 199 Klingenberg, M., 119, 126, 189, 190, 198, 297, 318,348 Klouwen, H., 137, 143, 144,193 Kobyashi, M., 167, 177, 198, 203, 336, 338,354 Kodicek, E., 161, 172,198 Kolahi-Zanovzi, M. A., 314, 315, 322, 349 Korda, F. H., 214,285 Korzybski, T., 142, I91 Kranz, Z. H., 193 Krebs, E. G., 337,354 Kristensen, B. I., 234, 282 Kroeger, H., 180, 198 Krogh, A., 119, 124,198 Kroon, D. B., 267,282 Kropf, R. B., 141,208,301,360 Kruyt, 274. Kubista, V., 292, 318, 326, 354, 356 Kuffler, S. W., 14, 23, 26, 28, 30 Kullenberg, B., 169, 198 Kumon,T., 39,67 Kurata, M., 89, 204 Kuroda, Y.,335,354

L Labows, J., 209, 211 Lafon, M., 117, 198 Lagwinska, E., 209,21 I Laidlaw, M., 275,281

367

Laidman, D. L., 166, 198 Lal, B., 118, 201 Lamb, K. P., 49, 50, 64 Lamberton, J. A., 193 Lambremont, E. N., 89, 92, 130, 189, 198,209,211 Lands, W. E. M., 142, 198 Langdon, R. C., 130,189 Langdon, R. G., 129, I98 Langernann, A., 210,210 Larner, J., 330, 331, 356 Laskowska, T., 345, 360 Lasser, N. L., 176, 192, 209, 210, 211 Laufer, H., 320, 353 Laurant-HubC, H., 311, 320,348,349 Law, J. H., 92, 168, 169, 181, 198, 207, 210,211 Lea, A. D., 178,206, 338,359 Leaback, D. H., 302,353 Leake, L. D., 12,29 Leclerqc, J., 13, 28 Lederer, E., 92, 93, 157, 168, 186, 188, 199,202 Lees, A. D., 255, 257, 260, 263, 271, 282 Legay, J. M., 160,199 Lehninger, A. L., 124, 139, 198, 199 Leibowitz, J., 290, 321, 354 Leifert, H., 37, 39, 40, 41, 64 Leloir, L. F., 307, 348 Lernonde, A., 117,206 Lenartowicz, E., 140, 141, 199, 346,354 Lennox, F. G., 39, 53, 64 Lester, R. L., 166, 199 Leto, S., 341, 343, 354 Levenbook, L., 99, 199, 291, 294, 326, 354 Levinson, Z. H., 150, 159, 160,161, 172, 173,198,199,204 Levy, H. R., 147, 176,199 Lewis, S. E., 139, 144,199, 202 L'Helias, C., 337, 354 Lindauer, M., 169, 190 Lingappa, B. T., 324, 358 Linzen, B., 169, 190 Lipke, H., 263, 275, 283, 341, 343, 354 Lippmann, E. 0.von, 291,354 Lisa, J. D., 37, 38, 39, 64 Littau, V. C., 278, 285

368

AUTHOR INDEX

Marchessault, R. H., 276,283 Marco, G., 263, 282, 344, 353 Marcuzzi, G., 294,355 Marsden, J., 350 Martin, A. J. P., 93, 200 Maruyama, K., 25,30 Maschwitz, U., 210,211 Maskell, E. J., 332, 352 Matsuhashi, M., 137, 200 Matsumura, S., 334, 355 Maurizio, A,, 295,298, 303,355 Maynard, L. A., 148,200 Mead, J. F., 130,200 Meinwald, J., 169, 191,209, 211 Melampy, R. M., 148,200 Meng, H. C., 113,193 Mercer, E. H., 222,283 Meyer, A. S., 183, 200 Meyer, H., 125, 200 Meyer, K. H., 214,219,283 Meyer, W. L., 135, 140,208, 346,360 Mezei, C., 209, 210 M Michalek, H., 167, 196 Micks, D. W., 43, 65 McAllan, J. W., 43,65 McCann, F. V., 6, 9, 10, 14, 20, 21, 22, Migicorsky, B. B., 176,200 Millard, A., 214, 284 28, 30, 31 Mishikata, T., 148, 200 McCarthy, R., 337, 338, 339, 355, 356 Mitchell, H. K., 37, 67,207 MacCay, C. M., 148,200 Mitlin, N., 43, 51, 65 McCloskey, J. A,, 169,198 Mitscherlich, 290, 355 McEnroe, W. D., 40, 65 Mittler, T. E., 43, 49, 65 McFarlane, J. E., 147, 200 Miura, K., 209, 211 Machne, X., 9, 10, 20, 28 Miyauchi, K., 174, 201 McInnes, A. G., 214, 283 Mochnacka, I., 42, 65, 289, 291, 293, Maciuca, C., 89, 200 296, 299,352,355 McKellar, J. W., 275,286, 345,359 Monod, J., 309,355 McLachlan, J., 214,283 Monro, A., 169,191 McLean, P., 147,200 Monroe, R. E., 131, 159, 161, 167, 171, McLintock, J., 267,283 172, 175, 179,197,199,201,202 McShan, W. H., 131,132,183,203,208, 307, 310, 311, 314, 315, 316, 320,360 Moore, B. P., 169,201 Moran, M. R., 117,201 Maddrell, 260,269,283 Morehead, F. F., 276,283 Maddrell, S. H. P., 260, 269, 283 Morehouse, M. G., 328,350 Maeda, M., 341,355 Morton, R. A., 166, I98 Maignon, F., 86,206, 359 Mosettig, E., 160, 172, 205 Malek, S. R. A., 223, 283 Moulinier, C., 327, 355 Maloeuf, N. S. R., 267, 283 Moyer, D. B., 153,199 Maltais, J. B., 49, 64 Miihlethaler, K., 214, 281, 283 Manchester, K. L., 184, 200 Mukaiyama, F., 41, 56, 64, 311,355 Manunta, C., 39,64, 148,200

Liu,F.,293,311,312,320,323,324,327, 348,355 Locke, M., 155,199,214,223,224,226, 251, 262, 263, 264, 265, 268, 278,281, 283, 341,349,355 Loeven, W. A., 267,282 Loomans, M. E., 137,194 Lotmar, 271 Loughheed, T. C., 291,293,323,360 Louloudes, J. S., 171, 206 Louloudes, S. J., 131, 153, 160,161,167, 172,197,199,202,205 Lowe, M. E., 140, 195 Lowenstein, J. M., 132, 197 Ludwig, D., 37, 38, 39, 47, 62, 64, 327, 328,355 Luick, J. R., 147, 199 Lullman, H., 23,28 Lum, P. T. M., 148,206 Lynen, F., 137, 166, 199,200 Lynn, J. W. S., 100,200

A U T H O R INDEX 369 Munday, K. A., 125,201 Nowosielski, J. W., 99, 201, 292, 301, M u ,E. A., 125,201 355 Murphy, M. R. V., 43,65 Numa, S., 137,200 Murphy; T. A., 30i, 306,307,308, 309, Nunez, J. A., 260,269,284 331.355 0 Murray, K. E., 193 Myrback, K., 290,291,355 O’Brien, R. D., 316, 356 O’Connor, A. K., 316,356 Ohad, D., 214,284 N Olson, J. A., 157, 201, 209, 211 Nair, K. S. S., 140, 150, 201, 327, 328, Orkand, R. K., 23,30 Orr,C. W. M., 336,338,356 351 Osborne, D. J., 161,193 Naito, K., 160, 195 Osman, M. F. H., 92,96,203 Nakahara, M., 160,197 Nakanishi, K., 160, 197 Osman, M. F. N., 92, 96,193 Osske, G., 168, 175,204,206 Nakase, Y., 320,354 Nath, V., 118,201 P Nation, J. L., 46,49, 51, 56, 65 Padmore, J., 153, 155, 188 Naudet, M., 113,192 Nayar, J. K., 160,205 Pain, J., 92, 93, 168, 186, 202 Needham, D. M., 25,30, 327,355 Panek, A., 315,356 Pant, J. C., 145,202 Needham, J., 33, 65 Pant, N. C., 145, 160,201 Nelson, M., 38, 40, 65 Pantel, J., 267, 284 Nelson, V. E., 277,286 Paoletti, R., 157, 194 Nelson, W. A., 52, 65 Paretsky, D., 98, 188 Nelson, W. L., 125,201 Park, C. R., 113, 193 Nettles, W. C., 327, 355 Parzen, S. D., 37, 65 Neufeld. E.. 315. 347 Neville,’A. C., 8; 30, 215, 216, 217, 218, Passey, R., 291, 324, 329, 351 221,223,227, 229,230,233,234,235, Passoneau, J. V., 341,345,356 Patterson, E. K., 138, 202 236,237,238,239,240,243,244,245, Patton, R. L., 37, 39, 40, 46, 49, 51, 56, 246,247,248,249,250,252,253,264, 265,266,268, 271, 272,273,283,284 61,65,99, 145,201,203,292,301,355 Payne, D. W., 311,322,350 Neville, E., 42,43,64 Newburgh, R. W., 125, 143, 189, 196, Payne, N. M., 51,65 Pearincott, J. V., 96, 202 209,210 Pearse, A. G. E., 330, 333,352 Newsom, L. D., 90,190 Pepper, J. H., 153, 188 Nicholas, H. J., 157, 201 Perryman, N. C., 100,200 Nicolai, E., 214,284 Petek, F., 314, 315, 322, 349 Niedergerke, R., 24,30 Niemierko, S., 85,86, 101,140,141,142, Petrushka, E., 137, 202 Petryszyn, C., 291, 293, 296, 299, 312, 144,199,201 320,355,356 Niemierko, W., 89, 100, 101, 117,201 Pfeiffer, I. W., 336, 356 Nijkamp, H. J., 43, 67 Philips, 319, 356 Nishizawa, K., 341, 355 Noble-Nesbitt, J., 227, 264, 265, 278, Philips, A. H., 129,198 Picken, L. E. R., 220,223,263,264,266, 284 271,282,284 Noland, J. L., 159, 201 Piek, T., 156, 202 Nordlie, R. C., 301, 355

370

AUTHOR INDEX

Pierce, W. D., 290,356 Pierre, L. L., 39, 65 Pipa, R. L., 266, 271,285 Piper, S. H., 93, 191 Poisson, R., 47, 48, 65 Polacek, I., 292, 318, 356 Pollard, A., 93, 191 Pollard, E. C., 285 Polonovski, J., 117, 144, 187 Popjiik, G., 132, 147, 157, 161, 176, 199, 202 Porembska, Z., 42,65,67 Porter, C. A., 209, 211, 344, 356 Porter, J. W., 132,202 Powning, R. F., 41,44, 50, 56, 65 Prairie, R. L., 170,205 Pratt, D., 346, 350 Pravdina, N. F., 327, 356 Preiss, B., 125, 200 Preston, J. M., 214, 284 Preston, R. D., 215, 217, 281, 284 Price, G. M., 144,202 Pringle, J. W. S., 7, 25, 29, 30 Prosser, C. L., 33,66 Prota, C. D., 37,66 Pryor, M. G. M., 263,264,266,272,276, 284 Pugh, E. L., 127,206

Q

Quastel, J. H., 137, 202

R Radcliffe, A. H., 198 Rajalakshmi, S., 130, 202 Ralph, C. L., 337, 338, 339, 355, 356 Ramazzotto, L. J., 327, 355 Ramsay, J. A., 152, 202 Randall, D. D., 291, 292, 296, 298, 310, 311, 313, 314, 322, 350,356 Randall, J. T., 214, 278, 284 Ray, J. W., 138, 196 Razet, P., 38, 39, 40, 41, 45, 46, 47, 48, 50, 51, 52, 54, 55, 56, 57, 58, 59,65,66 Reed, R., 214,284 Reger, J. F., 6, 29 Reid, J. A., 159, 194 Reiser, R., 145, 146, 206 Reiss, I., 25, 31

Rembold, H., 92, 190 Remmert, L. F., 139,199 Reuter, E., 264, 284 Ribi, E., 214, 284 Richard, A. G., 288,340,356 Richards, A. G., 138,202,214,221,222, 226, 229, 252, 264, 265, 266,271,276, 280,284,285 Ricketts, J., 143, 190 Ridge, R. M. A. P., 6, 30 Rieder, S. V., 310, 313, 314, 353 Ripley, S. H., 17, 30 Rittenberg, D., 70, 204 Robbins, W. E., 131, 159, 160, 161, 167, 171, 172, 175, 179, 185, 193, 197,199, 202,203,205,206 Robertson, M., 292, 353 Robinson, W., 39, 53, 66 ROCCO, M. L., 37, 39, 66 Roche, J., 42, 63 Rock, G. C., 145,203 Rockstein, M., 326, 356 Roeder, K. D., 33, 66 Roller, H., 183, 203 Romney, V. E., 119,194 Roosheroe, L., 43, 67 Rosell-Perez, M., 330, 331, 356 Ross, D. J., 38, 47, 66 Roth, L. M., 47, 66 Rothstein, F., 85, 203 Rousell, P. G., 327, 356 Rudall, K. M., 216, 217, 218, 219, 220, 223, 263, 266, 267, 269, 271, 273, 275, 281,285,288, 340, 357 Rudman, D., 184,203 Rudzisz, B., 141, 199 Ruegg, J. C., 25, 29 Ruiz-Amil, M., 302, 357 Ruska, H., 17,28 Rutschky, C. W., 117, 188

S Sacktor, B., 110,119,191,203,288,297, 302, 317, 318, 329, 333,357 Sahai, P. N., 93, 191 Saito, M., 167, 174, 177, 198, 201, 203 Saito, S., 293, 299, 300, 306, 312, 313, 326,357 Salpeter, M. M., 316, 356

A U T H O R INDEX Salt, R. W., 140, 203, 346, 357 Sarma, D. S. R., 130, 202 Sarma, P. S., 130, 202 Sauer, F., 127, 206 Savary, P., 98, 115, 116,192 Saverance, P., 37, 64 Scaria, K. S., 111, 116,194 Schaefer, C. H., 161, 175,203 Schild, H. O., 24, 28 Schildnecht, H., 210,210 Schlottke, E., 98, 203 Schmialek, P., 43, 63, 180, 203 Schmidt, E. L., 265, 278,285 Schmidt, G. H., 92, 96, 193, 203, 208, 211 Schmidt, W. J., 221, 229, 285 Schnatz, J. D., 115, 203 Schneiderman, H. A., 70,81,83,86, 119, 135, 140, 176, 178, 180, 181, 183, 194, 195,200,204,207 Schoenheimer, R., 70,204 Scholefield, P. G., 137, 202 Schrader, R. M., 89,198 Schreiber, K., 168,204 Schreiver, K., 175,206 Schroeffer, G. J., 130,204 Schulze, F. E., 224, 227,285 Schweizer, E., 137,200 Scoggin, J. K., 70, 89, 204 Scudamore, H. H., 234,276,285 Sedee, J. W., 53,66 Sedee, P. D. J. W., 43,67 Sedee, P. D. S. W., 131, 161,204 Sekhon, S. S., 225, 226, 285 Sembdner, G., 168,204 Senior, J. R., 98, 204 Senser, M., 291, 353 Setlow, R. B., 285 Setterfield, G., 266, 285 Shapira, A., 12, 29, 30 Shaw, D. R. D., 140,206 Shaw, J., 34, 51, 66, 67 Shearer, D. A,, 169, 189 Shigematsu, H., 302, 326, 330, 332, 356 Shikata, M., 93, 204 Shimizu, M., 174,203 Shortino, T. J., 159, 171, 172, 175,193, 197,202,203 Shyamala, M. R., 298, 306, 357

371

Siakotos, A. N., 103,204,263,275,283, 341,354, 357 Siddall, J. B., 210, 211 Siewerdt, R., 210, 211 Silverman, P. H., 150, 159, 199, 204 Silvestri, F., 227, 285 Sinha, R. N., 51, 63 Sinoda, O., 89, 204 Sipal, Z., 168, 193 Slhma, K., 183, 204, 338, 353 Slater, E. C., 119, 206 Slauttback, D. B., 143, 144, 193 Slepecky, R., 93, 204 Slifer, E. H., 117, 204, 225, 226, 285 Smissman, E. E., 173 Smith, A. F., 220,266, 280 Smith, D., 315, 316,357 Smith, D. S., 278,285 Smith, F., 346, 350 Smith, K. D., 37,66 Smith, M. J. H., 148, 196, 306, 352 Smittle, B. J., 155, 187 Smolin, A. N., 293, 296, 306, 327, 350, 356,357,358 Smyth, T. Jr., 209, 211 Sols, A., 303, 358 Somme, L., 346, 358 Souza, N. O., 315,356 Spiegler, P. E., 51, 66 Spiro, D., 301, 359 Spitteller, G., 179, I98 Squire, F. A., 140,204 Sridhara, S., 122,131,142,167,174,204, 205 Srivastava, P. H., 327, 358 Srivastava, P. N., 47, 56, 67 Staddon, B. W., 42, 46, 47, 51, 67 Stahler, N., 53, 67 Stamm, D., 186, 190 Staple, E., 157,205 Starkey, J. H., 153,199 Staudenmeyer, T., 303, 358 Stay, B., 326, 358 Steele, J. A., 160, 172, 205 Steele, J. E., 333, 337, 339, 358 Stein, C. I., 209, 211 Stein, G., 180, 205 Steinberg, D., 184, 206 Steinfelder, K., 175, 206

372

AUTHOR I N D E X

Todd, M. E., 292,359 Sten-Knudsen, O., 23,30 Tombropoulos, E. G., 128, 147,205 Stetten, Dew., 326,358 Tomita, M.,39, 67 Stetten, M.R.,326,358 Touter, O.,140,206 Steven, D.M.,255,285 Trager, W., 303,359 Stevenson, E.,326,333, 339,340,358 Stewart, C.M.,346,350 Travis, D. F., 227,243, 276,286 Treherne, J. E.,23,30,97,99,206,292, Stobbart, R.H.,34,67 297,298,304,359 Streibel, H.,296, 360 Strickland, K.P.,71, 135, 205 Trivelloni, J. C.,306,330, 359 Strong, F. E.,89,91,92,118, 130,205 Truszkowski, R.,37, 67 Tiimmler, R.,175,206 Strong, F. M.,137, 146, 191,194 Turner, R. B., 155, I87 Strumwasser, F.,262, 285 Subramanian, R. V. R., 275,281 U Sussman, A. S.,315, 324,352,358 Swann, M.M.,263,264,266,284 Uebel, E. G., 210,210 Sweeney, B. A., 239,285 Ursprung, H.,37, 39,66, 67 Synge, R. L. M., 93,200 Usherwood, P.N.R., 5,8,9,11, 12,13, Szarkowska, L.,42, 67, 167, 196, 312, 14, 15, 16, 17, 18, 20,27,27,28,30 320,356 Uvarov, B. P., 148,206 Szent-Gyorgyi, A., 266,285

V

T

Vallyathan, N. V., 11 1, 116,194 Tabor, L. A., 171, 175, 179,197,202 Vandenberg, J., 320,354 Tahmisian, T. N.,110,188 Van den Bergh, S. G., 119,206 Takata, N.,96,195 Vandenheuvel, W. J. A., 171,206 Takehara, I., 332, 346,358 Van der Kloot, W. G., 6, 19, 23,31 Takeshita, H., 326,357 Vanderzant, E. S., 145, 146,206 Takeuchi, A., 11,30 Vaney, C.,86,206-359 Takeuchi, N.,11,30 Van Handel, E.,148,178,206,209,211 Talalay, P., 157, 170,205 338,359 Tam, N.D., 186,190 Vardanis, A., 331, 359 Tanno, K., 296,346,347,358 Vaughan, M.,184,206 Tauber, 0.E.,70,89,204 Vaughan Williams, E. M., 26,30 Tauc, L.,262,280 Veerkamp, T. A.,267,282 Taylor, R. L.,226,264,276,285 Verhoeff, K. W., 264,286 Terahara, A., 160,197 Vickers, D. H., 43,51, 65 Terner, J. Y.,301,359 Vignon, P.,263,286 Terzian, L.A.,52,53, 64,67 Villeneuve, J. L.,117,206 Thompson, M. J., 154, 160, 171, 172, Von Ardenne, M.,175,206 180,190,197,203,205,210,2I0 Vonk, H.J., 43,67,209,211 Thomsen, E., 338,359 Voss, G., 311, 350 Thor, C.J. B., 218,219,286 Vroman, H.,153, 155,188 Thorsteinson, A. J., 160,205 Vroman, H.El, 167,171,175, 185,197, Tichomirov, A., 117,205 206 Tietz, A., 103, 131, 132, 133,202,205 Tietz-Devir, A., 70,99, 132,205 W Timon-David, J., 86, 148,205 Wagner, A. F., 137, 145,166,206 Timoshenko, S.,232,268,286 Wagner, R. P., 37,67 Tixier, M.,42, 63 Wakil, S. J., 127, 128, 132,202,206

373

AUTHOR INDE X

Waldbauer, G. P., 160,206 Waldvogel, G., 210,210 Walker, A. B., 209,210 Walker, R. J., 12, 29, 30 Walter, N. M., 276, 283 Wang, L., 263,282, 344,353 Watanabe, A., 3, 28 Watanabe, T., 160,207 Waterhouse, D. F., 56,67, 155,207,275, 286, 297, 320, 345,359 Waters, J. A., 160, 172, 205 Weaver, N., 92, 168, 207 Weber, A., 25,31 Weiland, G. S., 116, 193 Weinland, E., 148,207 Weinman, D., 296, 351 Weis-Fogh, T., 119, 124, 198, 207, 215, 216, 218, 219, 223, 243, 246, 267, 271, 272,280,281,282,286, 319,359 Weiss, E., 296, 360 Weiss, S. B., 135, 207 Werman, R., 6, 22,31 Wiechert, R., 210, 210 Wiens, A. W., 337, 339,359 Wiggers, H. A. L., 290, 359 Wigglesworth, V. B., 33, 44, 48, 49, 67, 99, 111, 148, 156, 207, 260, 263, 286, 336,360 Wilhelm, R. C., 135, 140, 207 Williams, C. B., 86, 207 Williams, C. M., 180,181, 183,204,207, 210,211, 340, 341, 345,352,356 Williams, E. F., 93, 191 Williams, R. H., 115,203 Wilson, E. O., 169,198 Wilson, G. S., 39, 66 Winston, P. W., 277,286 Wisniewska, A., 140, 141, 142, 168, 207 Wiss, O., 166, 195 Wlodawer, P.,85,86,100, 101,137,140, 141, 142, 144, 168,201,207,209,211 Wojtczak, A. F., 85, 86, 142, 144,201 Wojtczak, L., 137, 207 Wolfe, L. S., 137, 138, 207 Wood, D. W., 2,3,4,5,6, tl,12,13,14, 22, 23, 31 Woodring, J. P.,92, 189

Wootton, J. M., 161, 207 Wormser-Shavit, E., 297, 318, 357 Wren, J. J., 207 Wright, L. D., 157, 161, 176, 207, 208 Wright, T. R. F., 37, 66 Wyatt, G. R., 99, 135,140,141, 142,150, 188, 190, 208,290,291, 293,294,296, 299,300,301,306,307, 308,309, 310, 313, 314, 315, 316, 319, 323,326,327, 328, 329, 331, 333, 339, 340, 341, 342, 344,346,347,348,351,352,355,358, 360 Wyatt, S. S., 291, 293, 310, 313, 314, 315, 316, 319, 323,351,360 Wyckoff, R. W. G., 214,281 Wyman, J., 309, 355 Wyss-Huber, V. M., 296, 359

Y Yamafuji, I., 98, 208 Yamafuji, K., 86, 208 Yamaguchi, T., 341,355 Yamamoto, R. T., 180, 208 Yamashita, O., 296, 325, 332, 339, 340, 352,360 Yamazaki, H., 341,355 Yamazaki, M., 167, 174,203 Yonezawa, Y., 98,208 Yoshitaka, N., 56, 67 Young, R. C., 101, 156,208 Yuan, c.,210,211 Yushima, T., 311,360

Z Zahn, A., 43,63 Zaluska, H., 327, 328, 329, 341, 360 Zandee, D. I., 161,208,209,211 Zandee, D. L., 43,67 Zappanko, A., 98,194 Zborowski, J., 137,207 Zebe, E., 119, 123, 124, 125, 131, 132, 190,208 Zebe,E. C., 307,310,311,314,315,316, 320.360 Zielenska, Z. M., 56, 67, 345, 360 Ziv, O., 315, 347 Zubova, V. A,, 86, 87,-192

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Subject Index A abdomen, nerve cord, 23 absorption chitin, 328, 341, 345 lipid, 97-102 sugar from gut, 297-299, 320 water in chitin orientation, 277 Acanthocyclops viridis, cuticular orientation, 225 Acanthomyops claviger, fatty acid content, 95 isoprenoid content, 169 accessory sex glands, role in uric acid storage, 47 Acentrocneme hesperiaris, lipid content, 75 Acer negundo, sterols in, 175 acetate conversion to lipid in fat body, 148 conversion to sterol, 160,161,167,184 in fatty acid synthesis, 130-133, 147 in TGL and PL synthesis, 135, 137 acetate- 14C, oxidation, effect of corpus cardiacum, 337 Acetobacter xylinum cefulose polymerization, 263 electrostatic orientation mechanism, 279 acetylcholine, effect on muscle excitatory response, 12, 13 effect on postsynaptic potential in frog muscle, 9,10 spontaneous release in quanta1 units, 15, 16 Acheta domestica allantoicase activity, 39 nitrogenous excretion, 46, 47 uricolytic enzymes, 47 ACP (see Acyl carrier protein) Acrida bicolor, lipid content, 78 Acronycta rumicis, lipid content, 76 Acta caia, PL in, 142 ~ ~ + A . I . P4.

ACTH, sensitivity of lipase, 184 Actias lung, lipid content, 77 actin, in muscle contraction, 24, 25 action potentials, crural nerve trunk, 11 activation processes, in muscle fibre membrane, 22 active membrane response, in muscle, 8, 9 active transport in sugar absorption, 298, 322 muscle cell membrane chloride ions, 5 magnesium ions, 23 sodium extrusion mechanism, 6, 7 role of PL, 138 actomyosin, birefringence, 266 acyl carrier protein (ACP), in fatty acid synthesis, 127, 128 Acyrthosiphon pisum amino acids in honeydew, 49 fatty acid content, 94 glycogen and development, 327 Adelocephala heiligbrodtii lipid content, 77 adenase in ammonia formation, 42 in silkworm, 37, 41 in uricolytic pathway, 36, 37 adenine in uricolytic pathway, 34-36 oxidative deamination, 36 adenosine, enzymic deamination of, 36 adenosine deaminase in ammonia formation, 42 in uricolytic pathway, 36, 37 adenosine diphosphate (ADP) in carbohydrate metabolism, 305 in lipid metabolism, 69, 129, 136, 164 adenosine triphosphate (ATP) and nitrogen excretion, @O in carbohydrate metabolism, 305307, 319, 332

376

SUBJECT INDEX

adenosine triphosphate (ATPj c o n t . in lipid metabolism, 69, 102, 108, 109, 120-122, 129, 132-136, 138, 161, 164, 165 in muscle contraction, 24, 25

role in uric acid synthesis, 40 adipokinetic activity, 184, 185 adipose tissue, action of hormones, 184 ADP (see Adenosine diphosphate) adrenaline, effect on muscle excitatory response, 12 Aedes, glucose conversion to lipid, 148 Aedes aegypti fatty acid content, 95 nitrogenous excretion, 52, 53 nutrition and excretion, 53 PL in, 143 sterol utilization, 163 trehalose in eggs, 325 Aedes spp., neurosecretory cells and carbohydrate metabolism, 338 Aeschna see Aeshna Aeshna cyahea, nitrogenous excretion,

allantoic acid-cont. Orthoptera, 46 in uricolytic pathway, 35, 38 allantoicase in urea synthesis, 38, 39,41 in uricolytic pathway, 38, 39,45,47 allantoicotelic insects, excretory terminology, 59 allantoin end product nitrogen metabolism, 58 enzymic formation and degradation, 37-39

excretion Coleoptera, 50, 51 Diptera, 52-57 Hemiptera, 48, 49 Hymenoptera, 51, 52 Lepidoptera, 54-56 Neuroptera, 50, 51 Orthoptera, 46 in uricolytic pathway, 35, 37, 38 allantoinase in Collembola, 75 in Orthoptera, 47 in uricolytic pathway, 3 8 4 , 4 5 localization in insects, 40 allantoinotelic insects, excretory terminology, 59 allatectomy effect on carbohydrate metabolism,

46-48 Aeshna juncea, scalariform apodeme, 233 Aeshna sp., lipid content, 79 Aeshna spp., chitin orientation, 233, 234 age and lipid content, 81-85, 142, 169 336 effect on lipid metabolism, 180, 181, determination in cuticle, 245, 246 Aglais urticae, nitrogenous excretion, 184,185 54-56 allosteric, property of metabolic &ria afinis, fatty acid content, 93, 95 enzymes, 308, 309 Agritos comes, nitrogenous excretion, 5 5 ' ' amino acid oxidases in ammonia synthesis, 43 Alabama argillacea, lipid content, 75 albumin, and lipid, 102, 103, 106 in fatbody, 43 aldehydes, in cuticular wax, 153, 154 amino acids algae, trehalose in, 291 deamination, 42, 43 allantoic acid excretion end product nitrogen metabolism, 58 Coleoptera, 50, 51 enzymic formation and degradation, general aspects, 34,43,44 38, 39 haematophagous Diptera, 52, 53, excretion 57,58 Coleoptera, 50, 51 Hemiptera, 43, 44, 48-50 Diptera, 52-54 Homoptera, 49, 50 Hymenoptera, 51, 52 Orthoptera, 46 Lepidoptera, 54, 55 faecal material, 44

SUBJECT INDEX

amino acids-cont. in ammonia synthesis, 42, 43 in glutamic acid cycle, 43 in honeydew, 49 in purine synthesis, 34 in uricotelic pathway, 35 role in lipid metabolism, 137, 140, 147, 148 source of urea, 42 c-amino caproic acid, microfibril diameter, 214 4-amino-5-imidazole carboxamide riboside, role in uric acid synthesis, 40 ammonia biosynthesis amino acid deamination, 42, 43, 57 glutamic acid cycle, 43, 57 peptone deamination, 43 protein sources, 42,43 purine deamination, 42, 57 uricolytic pathway, 35, 38, 39, 42 end product of protein metabolism, 47,48 excretion aquatic insects, 42, 5I , 57 Coleoptera, 50,51 Diptera, 52-54 meat-eating maggots, 39, 42, 53, 54 minor product, 42 Neuroptera, 51 Odonata, 46-48 Orthoptera, 46 toxic end product, 34 role in uric acid synthesis, 40 ammonium bicarbonate, excretion, 51 Ammonotelic insects excretory terminology, 59 amylase in glycogen metabolism, 305, 334-336 pH in mid-gut, 320 Anabrus simplex cuticular lipid, 153 extra-cuticular hydrocarbons, 155 Anacridium aegyptium, lipid content, 78 androstenedione, 173 anisodiametric crystallites, parallel rearrangement, 265 Anopheles quadrimaculatis, nitrogenous excretion, 52, 53

377

Anopheles spp., lipid content, 80 antagonism of “relaxing factor” in muscle, 25 calcium-magnesium, in excitatory responses of muscle, 13, 14 Antheraea pernyi glycogen and development, 327 trehalose, 295, 296, 306 use of carbohydrate, 303 Antheraea spp. carbohydrate in larval hemolymph, 293 lipid content, 77, 86, 141 Anthomyiidae, lipid content, 79 Anthonomus grandis fatty acid synthesis, 130, 209 lipid content, 74, 89, 90, 92 nitrogenous excretion, 51 sterol in diet, 209 Anthophora sp., carbohydrate in larval haemolymph, 294 Anthrenus spp. glycogen and development, 327 Anthrenus vorax, lipid content, 140 aphid lipids in eggs of, 118 trehalase activity, 31 1, 319, 320 Aphididae, lipid content, 78, 91-94 Aphis brassicae, uricolytic enzymes, 50 .4phis fabae, trehalase activity in tissues, 311 Aphis pomi, fatty acid content, 94 Aphis spp., lipid content, 78 Aphrodite, X-ray diffraction of chaetae, 275 .4phrophora alni, chitin orientation, 234 Apis dorsata, lipid content, 81 Apis mellifera antenna1 cuticle structure, 222, 229 cuticular parabolic lamellae, 226 flight muscle and fatty acid oxidation, 125 and lipid hydrolysis, 111, 116 hexokinase activity, 302, 303 lipid content, 81, 93 metabolic oscillator, -252 monosaccharide utilization, 303 scent, 169

378

SUBJECT INDEX

Apis mellifera-cont. sugar in haemolymph, 294, 295, 298, 299 wax production, 156 Aplysia, circadian rhythms in neurons, 262 apodemes, chitin orientation, 220, 233 aquatic insects, excretion, 42,51,57 arabinose from plasma glycoprotein, 341 in Glossina spp., 296 Arachnida cuticle structure, 227 fatty acids in, 92 Arctia caja, lipid content, 74 Arctiidae, lipid content, 74 arginase in fat body extracts, 42 in ornithine synthesis, 42 in urea synthesis, 42, 57 arginine effect of arginase, 42 excretion, 35, 53 in ornithine cycle, 42 in urea synthesis, 42, 57 in uric acid synthesis, 40 use as phosphagen, 57 Argyrutaenia velutinana, fatty acids in diet, 145 Artemia salina chitin synthesis, 344 trehalose in, 324, 325 arthrodial membranes, chitin orientation, 223, 227 arthropodin chemical and mechanical properties, 217 chitin complex, 219, 269, 271, 272 fibre birefringence, 266 Ascaris carbohydrate synthesis, 329 trehalose in, 291, 324 aspartate, role in purine synthesis, 40,41 aspartic acid, effect on muscle excitatory response, 12 Astacus astacus, cuticle structure, 227 Asura conferta, lipid content, 75 ATP (see Adenosine triphosphate) Attacus spp., lipid content, 77

Attagenus piceus nitrogenous excretion, 50, 51 sterol utilization, 162 Aulacophorafumolaris, lipid content, 74 Automeris io, lipid content, 77 autoradiography in lipid metabolism studies, 156 of Calpodes cuticle, 263-265 of resilin lamellae, 234 Avena, cellulose reorientation, 266

B bacteria macromolecular orientation, 214 microfibril diameter, 214 Bacunculidae, lipid content, 79 Balaninus elephs, lipid content, 74 barium ions, effect on muscle fibre resting potential, 6 basalar muscle, peripheral inhibition of postsynaptic potentials, 18 bee behaviour and lipid content, 186 brain, 138 sex attractant, 180 solitary (see Ceratina) trehalose in diet, 319 bee larvae, fatty acids in, 92, 93 beetles, chitin orientation in cuticle, 220-222,226,232 behaviour and lipid content feeding, 185 mating, 86, 169, 186 worker bee, 186 Belostomatidae chitin orientation, 221, 231 cuticular structure, 229, 231, 232 Betulus alba, effect on Lasiocampa excretion, 55 Biblio marcia nitrogenous excretion, 52, 54 uricolytic enzymes, 54 biogenesis CoQ, 166, 167 sterol, 161, 164-167, 176, 209 biological clock mechanisms, in chitin orientation. 239

S UB J E CT I N D E X

biosynthesis ammonia, 35, 38, 39,42, 43, 57 chitin, 261, 262, 343, 344 CoQ, 166, 167 ecdysone, 179, 186 fatty acid, 127-134, 144, 146148 glucose, 301,302, 321 isoprenoid compounds, 161-1 69 lipid (see Lipid biosynthesis)

PL,134-137, 139-144 TGL, 134-137, 148 trehalose, 298, 304-309, 321

urea, 39,41,42, 49, 57 uric acid, 36, 37, 40, 41,47, 57 birds ornithine cycle, 42 uric acid synthesis, 40 birefringence artificial increase, 266 experimental control, 235 in experimental cuticle, 259 of cellulose, 220 of chitin, 220, 221, 235, 236 of constant-day cuticle, 238, 239 of drawn fibres, 266 of exocuticle, 235 of lobster cuticle, 221 in locust cuticle, 235, 236 biting actor, in silkworm diet, 160 Blaberus, muscle postsynaptic potential,

stuq 15, 16

Blaberus discoidali,s glycogen phosphorylase, 334 trehalase in tissue, 311, 313-315 Blaberus spp., endocrine system and carbohydrate metabolism, 338, 339 Blatta, &arthropodin in ootheca, 272 Blatta orentalis lipid content, 79 nitrogenous excretion, 46 Blattella, fatty acid synthesis, 131 Blattella germanica fatty acids in diet, 145 hydrocarbons in haemolymph, 155 lipid content, 79 sterol utilization, 159, 163, 171, 172

Blattidae lipid content, 79 14*

379

Blattidae-cont. uric acid storage, 47 Blissus leucopterus, lipid content, 78 blocking agents new0 muscular effect on electrical response, 24 effect on excitory response, 9, 11, 12, 13

effect on twitch tension, 24, 11 blood calcium content, 13 lipid in, 102, 103, 107, 110, 140 magnesium content, 13 blood sugar, regulation of, 299-301, 309,329

blowfly adenosine deaminase, 37 CoQ in larvae, 166 fat body lipid, 336 flight muscle and lipid function, 139 sterol modification, 172 uricase, 38 boll weevil fatty acids, 92, 130 lipid content, 74, 89, 90 bollworm, lipid in diet, 146 Bombus flight muscle, 18 sugar in hemolymph, 295 Bombycidae, lipid content, 74, 75 bombykol, as sex attractant, 186 Bombyx allantoicase activity, 39 and lipid metabolism, 117, 139, 144, 167, 174

and sterol biosynthesis, 167 and sterol modification, 174 diet, 160, 174 Bombyx mori amylase, 334 carbohydrate in haemolymph, 293 chitinolytic activity, 345 endocrine system and carbohydrate metabolism, 336, 338-340 glycogen metabolism, 326328, 332, 334, 341, 346

lipid content, 74, 75, 85, 86, 93, 96, 117,131,140,141,142,148

lipid in brain hormone, 177

380

SUBJECT INDEX

Bombyx mori-cont. monosaccharide utilization, 303 nitrogenous excretion, 56 pheronome, 186 phosphatases, 301, 302 sterols, 163, 168, 174 sugar absorption from gut, 298 trehalase activity, 311-313,322,323 trehalose, 296, 306, 325 bone collagen orientation, 221 parabolic lamellae, 223 bot fly (see Gastrophilus) Botrychium, trehalose in, 291 Bracon cephi, glycerol, 140, 346 brain function of PL, 138 hormone and lipids, 177, 178 effect on carbohydrate metabolism, 336,338,340 Brassica oleracea, 55 Brevicoryne brassicae, nitrogenous excretion, 49, 50 bromine, effect on amylase activity, 335 Bruchidae, lipid content, 73 Bryobia praetiosa active water balance, 217 bumble bee, sex attractant, 180 Buprestidae chitin orientation, 221 elytra structure, 232 lipid content, 73 butyrate, and fatty acid oxidation, 122, 125

C [14C]acetate,in lipid metabolism studies, 130,131,147,148,150,155,156,167 [l4C]glucose biosynthesis of trehalose, 304-307 blood sugar regulation, 299, 300 glycogen synthetase, 330, 331 lipid metabolism, 130, 132, 147-149 metabolism of chitin, 342, 343, 344 sugar absorption, 297,298 [l4CJgl&ose-1-phosphate, in trehalase synthesis study, 305

[14Clglutamate, in nitrogen excretion studies, 53 [14C]mevalonate, in sterol biosynthesis studies, 161, 168, 180, 181 [14CJpalmitate, and lipid metabolism studies, 103, 104, 106, 108, 117, 121, 123,124, 148,150,151, 185 [14C]pyruvate, in trehalose biosynthesis studies, 305 14C02,in fatty acid synthesisstudies, 133 cactus, sterol in, 169 Calaudra, exocuticle fibrogenesis, 264 calcification of crustacean gastroliths, 234 calcium, and lipase activity, 100, 113 calcium ions effecton muscle fibre resting potential, 6 effect on spontaneous miniature potentials in muscle, 15 importance in excitation-contraction coupling process in muscle, 24-26 role in electrically excitable response of muscle fibres, 22 calcium-magnesium antagonism, in excitatory responses of muscle, 13, 14 Calliphora absence of arginase, 42 carbohydrate content, 299 fatty acid synthesis, 131, 209 sterol, 161, 179 Calliyhora erythrocephala, carbohydrate metabolism, 338 Calliphora spp., lipid content, 79,80 Callobruchus,sterol utilization, 159, 163 Callobruchus chinensis, lipid content, 73 Callosamia promethea, lipid content, 77 Calopedes ethlius, wax secretion, 155 Calophasia Iunula, fatty acid content, 95 Calosaturnia mendocino, lipid content, 77 Calosoma calidum, fatty acid content, 94 Calotermes flavicollis, isoprenoid content, 168 Calpodes, cuticle autoradiography, 263-265 Calpodes ethluis chitin lamellogenesis. 251 cuticle structure, 228

SUBJECT INDEX

campesterol, 174, 175 Camponotus vagus, lipid content, 81 Campylenchie latipes, fatty acid content, 94 Cancer pagurus cuticular orientation, 225 Capnodis milliaris, trehalase activity, 311 Carabidae chitin orientation, 221 lipid content, 73 Carausius leg muscle effect of acetylcholine, 12 ion basis of electrically excitable responses, 22, 23 membrane potential, 2 magnesium in blood, 13 muscle potentials, 2, 5, 6, 16 Carausius morosus allantoinase activity, 47 nitrogenous excretion, 46, 47 carbohydrate content in haemolymph, 291-298 conversion to lipid, 86, 130, 132, 137, 147-149, 150, 151 carbohydrate levels, and diapause pupa, 300,342,345,346 carbohydrate metabolism effect of hormones, 309,336-340 effect on injury to diapause pupae, 333,339,340 general introduction, 287-289 interrelation of tissues in, 321 carbohydrates as flight energy source, 329 relation of chitin to, 328, 341-343 synthesis from fat, 328,329 Carbon dioxide effect on muscle potential, 6, 14, 26 role in purine synthesis, 40,41 Carcinus muscle flight muscle metabolism, 125 temperature and resting potentials, 6 Carcinus moenas, cuticular orientation, 225 Carex, 291

38 1

carnitine, in lipid metabolism, 125, 126, 143 /?-carotene, in dermal light sense, 255 carotenoid metabolism, temperature coefficient, 255, 256 Carpocapsa pomenella, lipid content, 76 Carposina niponensis, lipid content, 75 catabolism ecdysone, 186 fatty acid in embryos, 116-1 18 in flight muscle, 118-1 27 cations, divalent (see divalent) CDP (see Cytidine diphosphate) Cebrio gigas, lipid content, 73 Cecidomyiidae, lipid in, 209 Celerio euphorbiae carbohydrate in haemolymph,293,299 ornithine cycle, 42 PL in, 142, 143 trehalase activity, 312, 320 trehalose in haemolymph, 291, 296, 297 cell phospholipids, 137-139 trehalase in muscle, 314-316 cell membrane and sterols, 176, 178, 180 muscle excitation, 1, 19-23 phospholipids, 137, 138, 144 cell wall cellulose orientation, 215, 263 differentiation, 213 cellobiose, as inhibitor in trehalose synthesis, 309 in locust haemolymph, 292,295 cellulose birefringence, 220 comparison with chitin and tonicin, 213, 214, 227 crystallite orientation, 266 microfibril diameter, 214 orientation, 215 polymerization, 263 role of water in crystallization, 278 Cerambyx scopolii, lipid content, 73 Ceratina spp., sugar levels and overwintering, 296 Cetonia aurara, lipid content, 74

382

SUBJECT INDEX

Chaetocarabus intricatus, nitrogenous excretion, 50 Chalcophora mariana, carbohydrate in larval haemolymph, 294 Chilo simplex, trehalase activity, 311 Chironomusspp., and lipids, 80, 180 Chironomus thummi, trehalase activity, 311,320 chitin absorption, 328,341, 345 anisotropic character, 217 as a carbohydrate reserve, 328, 341 as a glycoprotein, 340 birefringence, 220, 221, 235, 236, 238, 239,266 chromatography, 236,237 comparison with cellulose and tonicin, 213, 214, 227 content of cuticle, 238 content of rubber-like cuticle, 248 1-4t!? covalent linkage, 215,217 crystallization, 215 effect of water on tensile strength, 218 fibril formation, 215 growth rate, 250 H-bonding, 215,217,218 histochemical localization, 236 in various body parts, 220,222 inhibition of lamellogenesis, 236, 238, 239 lamellogenesis rhythm, 249-252 levels of acetylation, 275, 276 macromolecular configuration, 217 metabolism and mounting, 328, 341-345 enzymatic degradation, 344, 345 general, 288 relation to other carbohydrates, 328,341-343 synthesis, 323, 329, 343, 344 microfibril diameter, 214 molecular transport, 215 orientation changes, 214,215 orientation in arthrodial membrane, 223 orientation in cuticle, 213-279 and pH, 274-277 anisotropic skeletal strains, 266 around tactile pits, 229,231,232

chitin-cont. association with protein, 269, 271. 272 biological clock control, 239 cellular regulation, 272 chemical bonds, 217,218 chemical control, 254, 260, 262 circadian clock control, 221, 222, 233-246, 259 conclusion, 279 control mechanisms, 215, 233-262 crossed fibrillar, 214, 220-223 cuticulin expansion, 223 daily rhythm systems, 233, 234 diurnal and nocturnal, 236 during intermoult growth, 268, 269 during moult, 267,268 exocuticular lamellae, 235, 236 experimental conditions, 238. 239, 269 extracellular polymerization, 263, 264 facultative lamellogenesis, 243, 244 form birefringence studies, 220, 235 functional aspects, 229-233 gel rearrangement, 265 Hemiptera, 221, 229, 231, 232, 234 hypotheses, 261,262,272-279 implantation experiments, 257-260 in deposition zone, 272 in relation to hardening, 267 in various arthropods and tunicates, 221, 222, 225-227,229,231,232, 239 independent of synthesis, 238 infra-red spectroscopy, 217 initiation, 251 internal and external factory, 261, 262 ionic changes, 276,277 ionic concentration, 272, 274, 275 lamellar structure, 223-229, 235238 mechanical properties, 217-220 mechanisms, 262-279 metabolic switches, 253, 254 Moir6 effect, 225, 228, 229 muscular forces, 267

SUBJECT INDEX

chitin-cont. nervous control, 260 obligatory lamellogenesis, 243, 244 of apodemes, 232,233, 267 of bristles and scales, 222, 263 of cross bars, 232 of egg shell, 226 of elastic ligament, 223 of elytra, 232 of exuvia, 226 of locust hind tibia, 237, 240 of pore canals, 236, 238 of prealar arm, 246-249 of puparium, 223,267 of tracheal taenidia, 222 of wing hinge ligament, 253, 254 organic and inorganic, 214 orthogonal fibril systems, 235 parabolic lamellae, 223-229 parallel orientation, 220 polarization analysis, 217 primary, 215, 262-265, 269 rhythmic lamellogenesis, 246-249 rhythmical water absorption, 277 scheme of events, 215 Schmidt's layer, 265 secondary, 215,262, 265-269 structural stability, 229-233 temperature effect, 239, 241, 242, 244, 245 types of, 220-223 uncoupling of clock, 241-243 X-ray diffraction study, 217 orientation mechanisms, 262, 263 oscillating synthesis, 246-254 synthesis pathway, 261 tensile strength, 217-219 van der Waal's forces, 217, 218 a-chitin molecular structure, 216 chitinase, 275, 344, 345 chitinolytic activity, in tissues, 345 chitobiase, 344, 345 chitogenetic cells, fine structure, 229 chitosan, in rubber-like cuticle, 246 chitosan-iodide, staining of exocuticle. 235, 238 chloride ions active transport of, 5

383

chloride ions-cont. effect on muscle fibre membrane, 4, 5, 20 effect on muscle inhibitory postsynaptic potential, 19 effect on muscle resting potential, 2,4, 5

chlorine, effect on amylase activity, 335 cholestanol, in lipid metabolism, 158, 162, 163, 177 cholestanone, structure, 158 cholesteine, structure, 158 cholestenone (ol), as growth factor, 162, 163 cholesterol, in lipid metabolism, 157169, 171-179, 184 choline, effect on muscle resting potential, 5 chorion, 85 chromatography absorption in juvenile hormone studies, 183 column in lipid studies, 90, 104, 105, 153, 168, 182 gas

in juvenile hormone studies, 181183 in lipid studies, 91-93, 145, 146, 153,155,169,177,178,186 infra-red in lipid studies, 172, 177 of chitin, 236-238 paper, in lipid studies, 92 in sugar analysis, 294-296, 304 in trehalase study, 313 silicic acid for lipid, 89 for juvenile hormone, 181, 183 thin layer in juvenile hormone studies, 183 ultra-violet in lipid studies, 172 chromosomes, parabolic lamellae, 223 chrysalis oil, 168 Chrysobothris ufinis nitrogen metabolism,, 50 Chrysomelidae, lipid content, 73, 74 Chrysopa curneu, uric acid storage, 51

384

SUBJECT INDEX

chylomicron, and lipid transport, 103, 110 Cicada, eye lens cuticle lamellogenesis, 264 Cicadidae, fatty acid content, 94 circadian clock definition, 239 graded uncoupling, 254,255,257 in chitin orientation control, 221,222, 233-246,259 in cuticle lamellogenesis, 233-246, 254,255 in homeostatic mechanisms, 244 in nervous system, 262 photoperiodic initiation, 254-257 temperature coefficient, 239, 244, 245 uncoupling experiments, 239-243 circadian rhythm, and blood trehalose regulation, 292, 300, 301 Cistelidae, lipid content, 74 Citheronia regalis, lipid content, 75 citrate and amylase activity, 335 and trehalase activity, 322 citric acid cycle, in liquid metabolism, 121, 122,124,125,132, 148 citrulline, precursor of urea, 42 classification, lipid, 71, 72 cleavage, and use of trehalose, 309-3 16, 321 CoA and CoASH (see Coenzyme A) Coccus cacti, lipid content, 78 cockchafers (see Melolonthu) cockroach allatectomy, 184 corpus cardiacum and carbohydrate metabolism, 337 lipid utilization, 106-108, 113, 125 muscle excitatory response, 12, 14 mechanical properties, 27 odour, 97 sterol modification, 171, 172 uricolytic enzymes, 37 Cocoon spinning, and lipid content, 85, 86 coenzyme A (CoA) in chitin synthesis, 261

coenzyme A (&A)-cont. in lipid metabolism, 69,120-123, 126129,132-134,136,161,164,165,184 coenzymeASH(CoASH),in lipidmetabolism, 69,120, 126, 129, 136, 164 coenzyme Q (CoQ), in lipid metabolism, 69,139, 166, 167 cofactors, in lipid metabolism, 121, 122, 124, 132, 133, 139, 140 Colaspidema atrum, lipid content, 73 cold resistance, and glycerol and sorbitol production, 325, 346 Coleoptera fatty acid content, 94 lipid content, 73, 74 nitrogenous excretion, 50, 51 uric acid storage excretion, 51 uricolytic enzymes, 50, 51 collagen fibre orientation in bone, 221 lamellar structure, 223 molecular orientation, 214 Collembola cuticle structure, 227 nitrogenous excretion, 45, 46 uricolytic enzymes, 45 Colocasia coryli, lipid content, 76 Colorado potato beetle, isoprenoid content, 168 complex lipids, 72 compound lipids, 72 Compsilura concinnata, nitrogenous excretion, 52 Conistra vaccinii nitrogenous excretion, 55, 56 uricolytic enzymes, 56 copepods, chitin in spermatophore, 263, 264 copulation, role of uric acid, 47 CoQ (see Coenzyme Q) Corcyra cephalonica, nitrogenous excretion, 56 Cordyalus cornutus, fatty acid content, 94 cornea, collagen orientation, 223 corpostanol, as growth factor, 162, 163 corpus alIatum and carbohydrate metabolism, 336, 338

SUBJECT INDEX

corpus allatum-cont. and lipid metabolism, 177, 180, 181, 184, 185 corpus cardiacum, and carbohydrate metabolism, 333, 337, 339, 340 cortisone, in water beetle, 210 cotton, tensile strength, 219 coupling process, excitation-contraction, in muscle, 23-27 crab (see also Carcinus) crab muscle, glutamate and contraction, 12 creatin, excretion, 44 Cremastogaster scutellaris, lipid content, 81 cricket, lipid in, 99 Croesus septentrionalis, lipid content, 81 crop, trehalase activity, 31 1 crural nerve trunk, effect of blocking agents on action potentials, 11 Crustacea calcification of gastroliths, 234 chitin microfibrils, 214 orientation in cuticle, 225, 227 muscle contraction, 23 facilitation, 14 pharmacology of inhibition, 19, 20 sodium ions and resting potential, 5 Ctenolepisma sp., and sterol biosynthesis, 167 CTP (see Cytidine triphosphate) Culex pipiens lipid content, 80 nitrogenous excretion, 52, 53 Culex tarsalis, fatty acids, 93, 96 curare, effect on potentials in muscle, 16 Curculionidae, lipid content, 74 Cuterebra, puparium formation, 267 cuticle age determination, 245, 246 and chitin metabolism, 340-344 chitin orientation, 213-279 (see Chitin) chitin-protein complexes, 269, 271, 272 elastic properties, 215, 216 fine structure, 223,229,236 growth, 247-250

385

cuticle-cont. growth layers, 235-238 inorganic ion content, 276, 277 lamellar stabilization, 229, 230 lamellogenesis action spectrum, 255, 257 chemical control, 254, 260, 262 circadian clock, 233-246, 254, 255 daily rhythm, 233,246-249 effect of daylight, 238,239,254-257 effect of temperature, 239,241,242, 244,245,255 environmental factors, 242, 243 facultative coupling, 243, 244 hypothetical cycle, 277 implantation experiments, 257-260 in wing hinge ligament, 253, 254 independent of sythesis, 238 initiating stimuli, 251 light threshold, 254-256 metabolic oscillation, 246-252 nervous control, 260 obligatory coupling, 243, 244 rates, 251, 252 rhythmical ion pump, 276,277 study of, 246 temperature coefficient, 239, 244, 245 laminate texture, 223-229,235-238 lipid in, 93, 131, 152-155 muscle insertions, 245, 246 percentage chitin, 238 physical properties, 215, 216 pigmentation, 256, 257 resilin, 216, 234, 246-249, 252-254, 272,273 solid and rubber-like, 215, 216, 223 strengthening bars, 232,233 swelling, 218, 229, 230 water uptake, 277 cuticle hydration, epidermal control, 277,278 cuticular scales, chitin orientation, 222 cuticulin, in chitin orientation, 223 Cynthia papillosa, parabolic lamellae, 224,225, 227 cysteine and activity of glycogen phosphorylase, 333

386

S U B J E CT I N D E X

cysteine-cont. and fatty acid synthesis, 132 cystine content in keratin, 50, 56 excretion, 35, 50 cytidine cofactors, role of, 139, 140 cytidine diphosphate (CDP), in lipid biosynthesis, 134, 135 cytidine triphosphate (CTP), in lipid metabolism, 135 cytochrome, and lipid metabolism, 108, 124, 125, 139, 166

D Dactynotus ambrosiae, fatty acid content, 94 Danaus plexippus, lipid content, 75 Daseychaeta alpim, lipid content, 76 Debye-Huckel theory, in chitin-protein complex, 274 Decticus verrucivorus chitin orientation, 234 population age dynamics, 245 definition, lipid, 71, 72 dehydrocholesterol, in lipid metabolism, 158,162, 163, 171-173, 177 Deilephila elpenor, carbohydrate in pupal hemolymph, 293 Deilephila sp., lipid content, 77 denervation, locust muscle, 16, 17 depolarization of muscle fibre membranes and electrically excitable responses, 8, 9, 11, 20, 21 and excitor axons, 7-9 and spontaneous activity, 23 and stimulation frequency, 14 effect of carbon dioxide, 6 effect of chloride ions, 5 effect of glutamate, 12 effect of transmitter substance, 17 in excitation-contraction coupling process, 23,24 depression, and time-dependence of muscle postsynaptic potentials, 14 derived lipids, 72 dermal light sense, 254-257 Dermaptera chitin orientation, 134

Dermaptera-cont. nitrogenous excretion, 46,48 Dermestes lipid content, 74, 89 sterol utilization, 159, 162 desiccation, role of cuticular lipid, 152154 desmosterol, in biosynthesis of cholesterol, 165 detergent, effect on trehalase activity, 316 detoxication, of metabolic end products, 58 development, and lipid metabolism, 84, 85, 93-95, 111, 112, 115, 131, 144, 150, 157, 159, 174, 175, 178,209 developmental stage and glycogen, 292-294, 301,327,331, 333,335,342 and sugar content, 291-297,309,323, 324 and sugar metabolism, 301, 303, 311, 323 DGL (see Diglyceride) diapause and glycerol production, 345,346 and lipids, 82, 85, 89, 90, 116, 117, 146 eggs in, 293,325,340,345 pupae in and glycogen phosphorylase, 333 carbohydrate levels, 293, 297, 299, 300,325,342,345,346 injury and carbohydrate metabolism, 333, 339,340 lipid content, 342 Diapause hormone, effect on carbohydrate metabolism, 336,339,340 diatoms, chitin structure, 214 Dictyoptera, chitin orientation, 234 Dictyostelium mucoroides, trehalose in, 324 diet and fatty acids, 90, 97, 129, 130, 145147 and gut trehalase activity, 319 and sterols, 157, 159-163, 168, 169, 172-176, 179,209 biting factor in, 160

S U B J E CT I N D E X

387

diet-cont. Donacia, aquatic cocoon, 263, 264 effect on blood sugar, 291,292,295,299 Donnan equilibrium, 2 effect on excretion, 50,53, 55-57 dormancy, and properties of trehalose, lipid-free, 97, 148 324, 325 wax in, 100,101,140,141 Drosophila differentiation adenosine deaminase, 37 of cell walls, 213 arginine requirement, 42 of skeletons, 213 chitin orientation, 263 diffusion, passive in sugar absorption, lipids and diet, 209 298,299,320,321 ornithine cycle, 42 digestion xanthine dehydrogenase, 37,39 and amylase, 334, 335 Drosophila melanogaster chitin, 275,341,342,344,345 amylase in, 334, 335 CoQ as product of, 166,167 bristle strength, 271 extra-digestive lipases, 110-1 16 fatty acid content, 92,95 lipid, 97-102, 127 sterol utilization, 163 wax, 100, 101 Drosophila pachea, sterol in diet, 169 digestive tract (see Gut) Dysdercus fasciatus diglyceride (DGL), in lipid metabolism, nitrogenous excretion, 48 69, 97, 103-110, 113, 116, 118, 127, urea synthesis, 48,49 134,136,138,209 Dytiscus marginalis dinoflagellate chromosomes, parabolic carbohydrate in haemolymph, 294 lamellae, 223 lipid content, 74 diolein, hydrolysis, 113 Diplopoda, cuticle structure, 227 E Diprion hercyniae, carbohydrate in ficles imperialis, lipid content, 75 haemolymph, 294 Diptera ecdysial glands, 177 ecdysone fatty acids in, 93,95, 131 flight muscle, 123, 124 and lipids, 159, 171, 172, 177, 179, lipid content, 79, 80 180, 184, 186,210 nitrogenous excretion, 52-54 effect on carbohydrate metabolism, PL in, 142, 143 336, 338 urease activity, 39 Echinops persicus, 290 uricotelic excretion, 53, 54 ecology, and lipid to carbohydrate condisaccharides, absorption from gut, version, 152 299, 320 and metabolic specialization, 169 dityrosine EDTA in tesilin orientation, 234 (see Ethylenediamine tetra-acetate) divalent cations effect on muscle resting egg development, and lipids, 82, 84, 85, potential, 6, 25 87,96,99, 116-118 Dixippus egg shell, chitin orientation in cuticle, lipid metabolism, 117 226 sugar levels in haemolymph, 337 eggs Dixippus morosus, lipid content, 79 carbohydrate metabolism, 296, 325, Dociostaurus marcoccanus, lipid content, 335, 340, 345 78 lipids in, 74, 75, 78-83, 117, 118, 144, Dolichopoda linderi, chitin orientation, 172-175 234, 239 Elateridae, chitin orientation, 221 1s 4- A.I.P. 4

388

S U B J E C T INDEX

electrical excitability comparison of “fast” and “slow” muscle fibres, 26, 27 of muscle fibre membrane, 2623,8,9 electrical response, effect of neuromuscular blocking agents, 24 electron microscopy cuticle fine structure, 223, 229, 236 in lipid digestion studies, 98 of Collembolan integument, 264 of constant day cuticle, 239 of forming cuticle, 265 of rubber-like cuticle, 246 electrophoresis detection of amylase, 335 in separation of plasma proteins, 341 of haemolymph, 103, 104, 108, 109 elytra, chitin orientation in cuticle, 232 embryo fatty acid catabolism, 116-1 18 lipid in, 139, 140, 175, 209 glycogen conversion, 345, 346 Embryogenesis and glycogen metabolism, 327, 332, 345 role of lipids, 70, 85, 102, 117, 118, 144, 174, 175, 185,208,209 trehalose changes during, 296, 325 end plate potential (see Postsynaptic potential) end products, of nitrogenous excretion, 33-61 endocrine control of wax secretion, 155 over uric acid deposits, 47 endocrine system, and carbohydrate metabolism, 288, 309, 336-340 endocrinology, and lipids, 176-186 endocuticle and chitin metabolism, 341, 343, 344 chitin lamellogenesis, 249-252 chitinous parabolic lamellae, 226, 227 lamellate structure, 236-238 pure preparation, 259, 260 Q l o of growth, 244,245 environment and blood sugar levels, 299 and cuticle lamellogenesis, 242, 24,3 and fatty acids, 93, 96, 209

enzymes amino acid oxidases, 43 in carbohydrate metabolism effect of hormones, 336, 337, 339, 340 glycerol, 346 glycogen, 305,326,329-337,340 monosaccharide, 301-305 trehalase, 309-324 trehalose synthesis, 305-309, 337 in chitin metabolism, 261, 275, 343345 in lipid metabolism, 97-101, 116116, 120, 125-127, 141, 143, 161, 165167, 170, 173, 176, 184 of ammonia formation, 42, 43 of urea formation, 41, 42, 57 of uricolytic pathway, 36-40, 45, 47, 56, 57, 59 of uricotelic pathway, 40,41 enzymic assays, for carbohydrate levels, 289,294,296 Ephestia chitin orientation, 263 Ephestia figulilella, lipid content, 76 Ephestia kuhniella, fatty acids in diet, 145 sterol utilization, 162 epicholestanol, as growth factor, 162, 163 epidermis and carbohydrate metabolism, 321, 323 chitinolytic activity, 345 control of cuticle hydration, 277, 278 trehalase in, 312 epithelium, and fatty acid absorption, 98 Ergates faber, carbohydrate in haemolymph, 294 lipid content, 73 ergostanol as growth factor, 162, 163 in sterol modification, 173 ergosterol as growth factor, 162, 163 in brain hormone, 177 in sterol modification, 172, 174 structure, 158 Eriosomatidae, lipid content, 78

389

SUBJECT INDEX

Eristalis tenax, lipid content, 80 esterases, in lipid hydrolysis, 111,112,l I5 esters, in cuticular wax, 153, 154 Estigmene acraea, lipid content, 74 ethylenediamine tetra-acetate (EDTA), and muscle contraction, 24, 25 Euphorbia, 320 Euproctis chrysorrhoea, lipid content, 75 Eurycotis floridana fatty acid synthesis, 131 sterol in, 175, 176 Eurythyrea marginata, lipid content, 73 Euttetix tenellus, lipid content, 78, 89 Euxoa segetum, lipid content, 75 excitation of skeletal muscles (see Skeletal muscles) excitation-contraction coupling process, in skeletal muscle action of carbon dioxide, 26 calcium ions, 24-26 depolarization, 23, 24 “fast” and “slow” muscles, 26, 27 excitatory postsynaptic potentials effect of GABA, 19 effect of picrotoxin, 19 excitatory responses in neuromuscular transmission action of carbon dioxide, 14 blocking effects of tryptomine, etc., 11,12 calcium-magnesium antagonism, 13, 14 general nature, 8, 9 ionic basis of postsynaptic potentials, 9-11 nature of transmitter substance, 11, 12, 13 time-dependent properties, 14 excitor axons, in muscle innervation, 7,8 excretion and diet, 50, 53, 55-57 nitrogenous (see Nitrogenous excretion) urea (see Urea) uric acid (see Uric acid) exocuticle chitinous parabolic lamellae, 227 lamellar stabilization, 229, 230 lamellate structure, 235-236, 264, 265

exocuticle-cont. extracellular fluid, ionic distribution between cell and muscle, 2 extraction, lipid, 71-73 eye collagen orientation, 223 lens cuticle, 264

F facilitation, in neuromuscular transmission, 14, 18 FAD (see flavine adenine dinucleotide) farnesol and juvenile hormone activity, 180183 as sterol precursor, 161, 166, 176 “fast” and “slow” axons electrically excited responses, 20, 22 in excitatory response of muscle, 8-14 in muscle innervation, 7, 8 inhibitory responses, 17, 18, 20 “fast” and “slow” muscles, difference between, 26,27 fat as flight energy source, 329, 334 conversion to glycogen, 327, 328, 329 fat body allantoinase, 40 amino acid and oxidases, 43 arginase, 42 effect of hormones on, 184, 185, 336, 338, 339 glucose metabolism, 301, 302, 307, 309, 321 glutamic dehydrogenase, 43 glycogen levels, 299, 300, 326, 329, 335,339,340 glycogen metabolism and trehalose synthesis, 305,306,321 effect of hormones, 336-339 general, 329,330 phosphorylase, 332-334, 337, 340 synthetase, 330, 331 glycoproteins in, 341, 343 lipases in, 111-1 16 lipid content during metamorphosis, 208 measurement of metabolism, 330 monosaccharide utilization, 303

390

SUBJECT I N D E X

fat body-cont. release of lipid, 102-108, 110, 111, 117,119,209 role in lipid metabolism, 99, 102-108, 110-117, 119, 125, 126, 131, 132, 139-142, 146, 148, 156, 184, 185, 208-2 10 storage of lipid, 99, 102-106, 146, 148, 184,209,210 trehalase activity, 311, 312, 324 trehalose biosynthesis, 304-309, 321 content, 297 uric acid storage, 47, 51 uricase, 39,40 xanthine dehydrogenase, 39 fatty acid composition, in various species, 90-97 fatty acids and nutrition, 145-147,209 catabolism effect of hormones, 185 in embryos, 116-1 18 in flight muscle, 118-127 digestion and absorption, 98-102,115, 116 function, 89, 92, 93, 97, 145, 146 in classification of lipids, 72 in cuticular wax, 153, 154 oxidation, 337 synthesis general mechanism, 127-130 in insects, 128, 129, 130-134, 144, 146-149, 209 feeding activity and blood trehalase activity, 324 and blood trehalose regulation, 299, 301 and cuticle reabsorption, 341 and glycogen production, 328-330 FFA (see Free fatty acids) lirebrat (see Thermobia domestica) flagellin molecular orientation, 214 flavine adenine dinucleotide (FAD), in lipid metabolism, 69, 120, 121 flax, tensile strength, 219 fight and haemolymph trehalose level,. 292,. 294, 317-3 19

flight-cont. role of glycogen, 317-319, 329, 334 flight muscle calcium ions and contraction, 25 cuticle insertions, 245, 246 effect of carbon dioxide on excitation, 6, 14 innervation, 8 peripheral inhibition of postsynaptic potentials, 18 potassium ions and membrane potential, 3 respiration, 118-127 role of carbohydrate metabolism energy sources, 329,333 glucose, 302, 317-3 19 glycogen, 317-319,329,330,333 sugar levels, 297 trehalase, 311, 312,314-316, 318 trehalose, 317-3 19 role of lipids, 70, 86, 102, 110-113, 115-127, 139,142, 144 three types, 123, 124 Flora, symbiotic, 130 fluorescence analysis of resilin lamellae, 246,247,253 fore-gut, trehalase activity, 311, 312 Forficula auricularia chitin orientation, 234 nitrogenous excretion, 46,48 formate, role in uric acid synthesis, 40, 41 Formica polyctena fatty acid content, 96 lipid content, 208 Formicidae, lipid content, 81 fowl, lipids in, 70 free fatty acids (FFA), in lipid metabolism, 69, 89, 92, 96-103, 105-108, 110, 112, 114, 118, 126, 132, 133, 139, 150, 156, 208,209 freezing and glycerol and sorbitol, 325, 346 freezing and thawing effect on trehalase activity, 314, 316, 319 frog (see also Rana) frog muscle excitation of “fast” and “slow” fibres, 26, 27

S U B J E C T INDEX

frog muscle-cont. postsynaptic potentials, 9, 15 role of ions, 2, 5, 13, L7, 23, 24, 26 frost-resistance, and sugar levels, 296 fructomaltose, in bee haemolymph, 295 fructose in haemolymph, 291-296, 298, 299 intestinal absorption, 298 utilization of, 302, 303 fructose-6-phosphate, in chitin synthesis, 261 fucose, from insect glycoprotein, 341 fungi, trehalose in, 290, 291, 310

39 1

glands-contd. corpus allaturo, 177, 180, 181, 185, 336-338 corpus cardiacum, 333,337,339,340 mammary, 147 mandibular, 169, 186 pituitary, 185 prothoracic, 172, 177, 178 salivary, 180 Glossina morsitans, nitrogenous excretion, 52, 53 Glossina spp. lipid content, 80 sugar content, 296 glucagon, sensitivity of lipase, 184 G glucosamine GABA (see Gamma aminobutyric acid) from plasma glycoprotein, 341 galactosamine, from plasma glycoproin chitin, 261, 340, 343, 345 tein, 341 inhibition of trehalase, 313, 315 galactose phosphorylation, 302 from plasma glycoprotein, 341 glucosamine-6-phosphate, and activain haemolymph, 295 tion of glycogen synthetase, 331 use of, 303 glucose galactose-6-phosphate,and activation of and biosynthesis of trehalose, 298, glycogen synthetase, 33 1 304-307, 309, 321 Galerucella luteola, lipid content, 73 and flight energy, 317-319 Galleria, lipid metabolism, 145, 168, 209 biosynthesis and utilization, Galleria mellonella general, 321, 303 carbohydrate in haemolymph, 293 hexokinases, 302, 31)s cuticle structure, 226 phosphatases, 301, 302, 305, 322 effect of exposure to cold, 346 conversion to lipid, 130, 132, 147-149 lipid in, 76, 95, 141 effect of hormones on level, 336-339 mucopolysaccharide in, 341 from glycoprotein in plasma, 341 nitrogenous excretion, 56 in chitin synthesis, 261 wax digestion, 100, 101, 140, 141 intestinal absorption, 297, 298, 320gamma aminobutyric acid (GABA), 322 effect on responses in muscle, 19, 20 levels, Gastrophilus in flight muscle, 297, 318, 319 carbohydrates in, 291, 294, 326 in haemolymph, 291-296, 299, 317, lipid content, 80, 99 322-324 Gelechiidae, lipid content, 75 occurrence, 289 genes, effect of ecdysone, 180 glucose-1-phosphate,306, 310,318,331Geotrupes stercoralis, lipid content, 74 334 geraniol, in scent, 169, 176 glucosed-phosphate, 261,301,305, 307giant axons, squid, 21 309, 318, 331 “giant” miniature potentials, in aener- glutamate vated muscle, 16, 17 as excitatory transmitter substance, 12 in purine synthesis, 40, 41 slands accessory sex, 47 role in arginine synthesis, 53

392

SUBJECT INDEX

glutamic acid effect on muscle excitatory response, 12 in ammonia formation, 43 in chitin synthesis, 261 glutamic acid cycle, in ammonia formation, 43 glutamic dehydrogenase, in ammonia formation, 43 glutamine in chitin synthesis, 261 in purine synthesis, 40,41 glutathione, and fatty acid synthesis, 132, 133 glycerol, levels in diapause and development, 300, 325, 326 glycerol production, 325, 345, 346 glycine, conversion to lipid, 148 glycine, role in purine synthesis, 40,41 glycogen and chitin as reserve nutrients, 328, 341 changes during growth and metamorphosis, 300, 301, 325, 327-329, 333, 335, 342, 345,346 conversion to lipid, 147-149, 150, 151 extraction, 326 in hemolymph, 292-294,326,327 in insect flight, 317-319, 329, 334 in insects, 325-327 in trehalose biosynthesis, 305, 306, 337 metabolism of, amylase, 305, 334-336 effect of, hormones, 337-340 in fat body, see Fat body phosphorylase, 305, 326, 329, 330, 331-334 synthetase, 330, 331, 305 storage in crop, 318 and moulting, 327-329, 341,342 reserves and nutritional changes, 299, 306 glycoproteins in insects, 296, 340, 341 metabolism of chitin, 341-345 glyoxylate cycle, 150, 152, 329

glyoxylic acid, in uricolytic pathway, 38, 39 Coliathus, tensile strength of chitin, 219 Gomphocerus maculatus, chitin orientation, 234 gonad maturation, effect of queen substance, 186 gonadotrophic hormone and lipid metabolism, 177, 180, 184 grasshopper allatectomy, 184 lipid utilization, 106-108 PL in diet, 160 growth and monosaccharide utilization, 302, 303 cuticle, 235-238, 247-250 glycogen accumulation and conversion during, 301, 327-329, 335, 342 role of lipids, 70, 84, 85, 96,97, 101, 145-147, 157, 162, 163 growth factors, 162, 163 growth hormones, 70, 81, 177, 179 Gryllodes sigillatus, fatty acid and growth, 146, 147 Cryllus domesticus carbohydrate in haemolymph, 292, 300, 301 sterol utilization, 163 GSH, 107 guanase in uricolytic pathway, 36, 37 in uricotelic pathway, 41 role in ammonia synthesis, 42 guanine in uricolytic pathway, 34-36 in uricotelic pathway, 41 guanosine, enzymic deamination, 36, 37 gut active transport of water, 298 amylase in, 334, 335 chitinase activity, 345 lipase, 98-101, 111-1 13 sugar absorption, 297-299, 320 sugar levels, 296 trehalase, 310-316, 319, 320, 324 trehalose, 319-322

S U B J E C T INDEX

gut wall, and carbohydrate metabolism, 320, 321, 335

gypsy moth, sex attractant, 186

H in lipid studies, 171, 172, 179 haematin, excretion in tsetsefly, 44 haemolymph amylase in, 334, 335 and chitin metabolism, 343, 345 and trehalose-trehalase physiology, 311, 312, 317-324

biosynthesis of trehalose, 304, 305, 309

“glycogen” in, 292-294, 326, 327 hydrocarbons in, 155, 156 lipase in, 111, 112, 113 lipid in, 97, 99-109, 116, 126, 135, 140-142, 146, 168, 178,209

magnesium in, 23 potassium concentration and muscle membrane potential, 4 sodium in, 22 sugar levels effect of hormones, 336-340 in insects, 291-298 regulation, 297-301, 309, 329 urea content, 41 Hamaker curve of, [K] influence on interaction energy, 274 Harpalus spp. lipid content, 73,94 Haversian system, collagen orientation, 221

head tissues, trehalose biosynthesis, 306 heart, mammalian and fatty acid synthesis, 128 role of PL, 138, 144 Heliocopris colossus, chitin orientation, 221 Heliotaurus spp., lipid content, 74, 75 Hemichroa alni nitrogenous excretion, 52 Hemiptera chitin orientation, 221, 229, 231, 232, 234 lipid content, 78 nitrogenous excretion, 43, 44, 4850

uricolytic enzymes, 48

393

Hesperidae, lipid content, 75 Heteroptera, nitrogenous excretion, 48, 49

hexokinases, in glucose biosynthesis and utilization, 302, 305 hexosamine and chitin metabolism, 343 protein-bound in fat body, 341 hind-gut, trehalase activity, 311, 312 histamine, excretion, 49 histidine excretion, 35, 49, 53 in uric acid synthesis, 40 histochemistry in lipid metabolism studies, 118, 140, 148

in study of chitin, 236 hog intestine, trehalase, 315 Homarus sp., chitin fibril orientation, 225,227

homoestatic mechanisms, involvement of circadian clock, 244 Homoptera fatty acid content, 91, 93 nitrogenous excretion, 49, 50 uricolytic enzymes, 50 honeybee (see also Apis mellifea) honeycomb, utilization of, 100, 101 honeydew amino acid content, 49 trehalose in, 321, 322 Hordeum mutinum, effect on h i o c a m p a excretion, 55 hormones adrenaline, 12 and isoprenoid compounds, 176-186 brain, 177-178 juvenile, 177, 180-1 85 moulting, 171, 172, 177, 179, 180 queen substance, 186 sex attractants, 186 cortisone, 210 ecdysone, 336, 338 effect of lipids on, 146 effect on carbohydrate metabolism, 288, 309, 336-340

effect on enzymes, 143 effect on lipid metabolism, 184, 336339

394

SUBJECT INDEX

hormones-cont. effect on respiration, 337, 340 effect on wax production, 155 gonadotrophic, 177, 180, 184 growth, 70, 81, 177, 179 juvenile and carbohydrate metabolism, 336 and lipids, 81, 336 mode of action, 177, 184, 185 purification, 180-184 synthetic substances, 210 moulting, 171, 172, 177, 179, 180 precursors, 146, 176 purification, 177, 180 vertebrate and lipid metabolism, 184 housefly (see also Musca domestica) conversion of glucose to lipid, 148 hexokinase activity in flight muscle, 302 sterol in diet, 160, 161 Hyalophora spiracular muscle effect of carbon dioxide on membrane, 6 spontaneous activity of membrane, 23 Hyalophora cecropia carbohydrate metabolism and flight, 329 and haemolymph, 293, 297, 299, 300 glucose, 301 “injury factor”, 339, 340 interconversion, 148-151 trehalases, 313, 315 trehalose, 304-308, 325 chitin and glycogen synthesis, 328 chitin metabolism, 341, 342, 344, 345 fat body glycogen synthetase, 331 glycogen phosphorylase, 333, 334 -haemolymph, 297 fatty acid oxidation, 121-126 lipid content, 77, 81, 85, 90,96, 103105, 117, 140-142, 182 lipid utilization, 103-1 16

Hyalophora cecropia-cont. overwintering and sorbitol production, 346 respiratory quotient, 87, 88 sterol in and hormones, 178-185 biosynthesis, 161, 167, 168 function, 176 modification, 174, 175 Hyalophora euryalus, lipid content, 77 Hyalopterus pruni, lipid content, 78 hyaluronic acid, in insect tissues, 341 hydration, cuticle, 277, 278 hydrocarbons cuticular, 152-1 55 extra-cuticular, 155-1 57, 169 Hydrocyrius, flight muscle, 25 Hydrocyrius colombiae chitin orientation, 221. 231, 234 circadian clock, 234, 239 cuticular structure, 23 1, 232 hydrogen bonding, of chitin, 215, 217, 218 Hydrous piceus, carbohydrate in hemolymph, 294 hydroxycholesterol, structure, 158 5-hydroxytryptamine, as a neuromuscular blocking agent, 11, 16 Hylemya brassicae, fatty acid content, 95 Hylotrupes bajulus, sterol utilization, 163 Hymenoptera carbohydrate in haemolymph, 294 cuticle structure, 226 fatty acid content, 95 lipid content, 81 nitrogenous excretion, 51, 52 uricolytic enzymes, 52 hyperpolarization of muscle fibre membrane and electrically excitable response, 22 and inhibitor axons, 7, 19 and peripheral inhibition, 17, 18 effect of carbon dioxide, 6 effect of chloride ions, 5 effect of GABA, 19 effect of temperature, 7

SUBJECT INDEX

hyperpolarizing potentials, and inhibitory postsynaptic potentials, 18, 19 Hyphantria cunea, fatty acid content, 95 Hypoderma bovis, secondary chitin orientation, 267 hypoxanthine enzymatic oxidation, 37 in excreta, 52, 56 in protein metabolism, 41, 58 in uricolytic pathway, 35, 36 in uricotelic pathway, 35, 41 “hysteresis”, effect of carbon dioxide on muscle, 26

I identification, lipid, 72, 73 implantation, in cuticle lamellogenesis studies, 257-260 inactivation process, in muscle fibre membrane, 21,22 infra-red chromatography, in lipid studies, 172, 177 infra-red spectrometry, in cuticular wax studies, 153 infra-red spectroscopy in lipid studies, 186, 187 of chitin orientation, 217 inhibition and neuromuscular transmission, 1720 of lamellogenesis of chitin, 236, 238, 239 inhibitor flight muscle respiration, 122 metabolic, 7 wax hardening, 153 inhibitor axons, action in muscle, 7, 1720 inhibitors in glyceride synthesis, 108 in sterol synthesis, 176 in trehalose biosynthesis, 308, 309 lipase, 113, 115 of trehalase activity, 313,315,322-324 inhibitory responses in neuromuscular transmission inhibitory postsynaptic potentials, 18, 19

395

inhibitory responses-cont. interaction with excitatory responses, 20, 18 ion basis of postsynaptic potentials, 19,20 peripheral inhibition, 17, 18 pharmacology of inhibition, 19, 20 injury of diapause pupae integument effect on blood trehalose level, 299 effect on carbohydrate metabolism, 333, 339, 340 innervation, of muscle, 7, 8, 15 integument chitinase activity, 345 effect of injury to diapause pupae, 299, 333, 339, 340 trehalase activity, 312 interconversion, lipid and non-lipid, 147-152 intestine (see Gut) ion concentration, and adenosine triphosphate in muscle contraction, 24, 25 ionic aspects, chitin orientation in cuticle, 272, 274-277 ionic basis of electrical excitable responses of muscle fibre membrane, 21-23 of excitatory postsynaptic potentials in muscle, 9-11 of inhibitory postsynaptic potentials in muscle, 19, 20 ion pump, in cuticle lamellogenesis, 276,277 ips typographus, trehalase activity, 311 isoprenoid compounds and insect hormones, 176-186 biosynthesis, 161-168,209 content, 168-170 function, 175, 176 nutritional studies, 157-161 sterol modification, 170-175 Isoptera cuticle structure, 227 lipid content, 79 isotropism, of chitin, 221

396

SUBJECT I N D E X

J

Lasioderma sericorne, sterol utilization,

162 lauric acid, 94,95 leafhoppers, lipid content, 91 lectithin, and PL synthesis, 143 leg muscle and fatty acid oxidation, 125 glycogen metabolism, 330,333 inhibitory responses in neuromuscular transmission, 18-20 innervation, 7, 8, 17 membrane potential, 2 trehalase activity, 310, 311 trehalose biosynthesis, 306 Lepidoptera K cuticle structure, 226 keratin fatty acids, 95, 125, 131 cystine content, 50, 56 juvenile hormone in, 183 dietary effect on excretion, 50, 56, lipid content, 74, 75-78, 81-86, 89 57 membrane potentials of muscle fibres, Kethocerus indicus, pheromonic effect of 4 lipids, 186 nitrogenous excretion, 54-56 a-ketoglutarate, in fatty acid synthesis, uricolytic enzymes, 56 133 a-ketoglutaric acid, in ammonia forma- Leptinotarsa decemlineata, lipid content, 73,94 tion, 43 Lethocerus, flight muscle, 25 L Lethocerus spp., chitin orientation, 221 lamellogenesis, of cuticle (see Cuticle) Leucania separata Lampyridae, lipid content, 74 carbohydrate in larval hemolymph, lanosterol, in biosynthesis of cholesterol, 293 165 trehalase, 311, 324 Laphygma spp., lipid content, 75, 76 leucine, conversion to lipid, 148 Laria irresecta, lipid content, 73 Leucophaea maderae Lurinus nidificans, and trehalose, 290 carbohydrate metabolism larva carbohydrate in haemolymph, 292 amylase activity, 335 effect of hormones, 339 glycogen during growth, 327, 328 trehalase activity, 311, 320 lipid content, 73-87, 92-97, 99-101, trehalase characteristics, 314-316 1?1,140-143 trehalase biosynthesis, 306, 307 sugar content, 291-294 metabolism, 117, 118, 185 trehalase activity, 311, 312, 314, 320 ligament, chitin orientation, 223, 253, trehalose synthesis, 304-306 254 larval cuticle, lipid in, 154, 155 light larval diet and chitin orientation in cuticle, 238, and excretion, 55 239 and lipids, 145-147, 159, 160 and fatty acids, 96 Lusiocampa trifolii, larval diet and exdermal sense, 254-25'7 cretion, 55 effect on cuticle lamellogenesis, 238, Lasiocampidae, lipid content, 75 239,254-257

Jassidae, lipid content, 78 junction potential (see Postsynaptic potential) juvenile hormone effect on carbohydrate metabolism, 336 effect on lipids, 336 lipid content, 81 mode of action, 177, 184, 185 purification, 180-184 juvenile hormone activity, of synthetic substances, 210

SUBJECT I N D E X

397

Limogeton fieberi, chitin orientation, lipid content+ont. 221 isoprenoid compounds, 168-170 Limulus sp., cuticle structure, 227 of various species, 73-81,88 1-4 linkage, of chitin, 215,217 lipid metabolism effect of hormones, I inoleic acid 336-339 in lipid metabolism, 91, 92, 9497, lipid utilization 130,145 digestion and absorption, 97-102 in lipid metabolism, 91, 92, 94-96, extra-digestivelipases, 110-116 126,130 fatty acid catabolism, 107,116-127 lipase general mechanism, 185,203 111-113,115 release and transport, 102-1 11, 117, extra-digestive, 110-116 119,209 gut, 99-101,111-113 lipoprotein membranes, and trehalase sensitivity to hormones, 184 location, 316 b a s e activity, effect of calcium, 100,113 lipoproteins, 103, 108-1 10 lipid Lithosiidae, lipid content, 75 and diapause, 82, 85,89,90, 116, 117 liver and insect hormones, 176186 mammalian glycogen metabolism, 330, 332 classification, 71,72 sterol biosynthesis, 176 components, 72, 89 lobster, cuticular properties, 221 cuticular, 93, 131, 152-155 locust definition of, 71, 72 diet extracuticular, 155-157 and lipid content, 146 fatty acid composition, 72, 89-97 hydrocarbons and waxes, 152-157 effect of terpenes, 209 fat body isoprenoid compounds conversion to lipid in, 148 ‘biosynthesis, 161-168,209 content, 168-170 fatty acid synthesis, 131, 132, 133 flight muscle function, 175, 176 hormones, 176186 and fatty acid oxidation, 119, 124, nutrition, 157-161 125 sterol modification, 170-175 and lipid hydrolysis, 119 metabolism and function, 69-187 hexokinase activity, 302 method of extraction, 71-73 potassium ions and membrane nature of, 89-97 potential, 3 lipid biosynthesis haemolymph fatty acid biosynthesis, 127-134 carbohydrate in, 295 fatty acids in nutrition, 145-147 trehalase in, 322 PL and TGL, 134-144 leg, substrate interconversion, 147-152 chitin orientation in cuticle, 237, lipid content 240 alterations during metamorphosis, leg muscle 81-89 excitatory postsynaptic potential, and developmental stage, 73-86, 89, 10 93-95,118, 131 “fast” and “slow” fibres, 27 and sexual dimorphism, 81,8483.96, innervation, 7, 8, 17 monosaccharideutilization, 303 97,99 expression of, 71-73,84 muscle excitation, 12, 13, 16, 17,23,25 in diapause pupa, 342

398

SUBJECT INDEX

locust-cont. glycogen phosphorylase, 333 glycogen synthetase, 330 PL in, 141 spiracular muscle contraction, 24 trehalose biosynthesis, 306 Locusta leg muscle electrically excitable response, 22 membrane potential, 2,4 monosaccharide utilization, 303 muscle postsynaptic potential, 17-20 sodium ions and resting potential, 5 Locusta migratoria carbohydrate content, 292, 326 chitin orientation, 234 circadian clock, 234, 239 flight and carbohydrates, 297, 318 and fatty acids, 96, 126 lipid metabolism fatty acids, 96, 126, 133, 145 lipid content, 78, 79, 92, 99, 105, 117, 144 sterol utilization, 160, 162 nitrogenous excretion, 46 uncoupling lamellogenesis, 241 Locustapardalina, lipid content, 79 London-van der Waals forces, in chitin and protein orientation, 274 Loxostege spp., lipid content, 76, 77 Lucanidae, chitin orientation, 221 Lucilia creatin excretion, 44 urease, 39 uricase, 38, 54 Lucilia cuprina, ammonia production, 39 Lucilia sericata lipid content, 80 nitrogenous excretion, 52-54 sterol utilization, 157, 162 trehalase activity in tissues, 311 Luciola vitticollis, lipid content, 74 lung mammalian and fatty acid synthesis, 128 carbohydrate interconversion to lipid, 147

Lycophotia margaritosa, lipid content, 75 Lygaeidae, lipid content, 78 Lymantria dispar, lipid content, 75 Lytta vesicotoria, lipid content, 74

M Maciosiphium bari, lipid content, 89 Macropipus puber, cuticular orientation 225 magnesium and fatty acid oxidation, 121, 122, 124 and fatty acid synthesis, 132, 133 And haemolymph trehalose activity, 322 in trehalose biosynthesis, 308, 309 magnesium-calcium antagonism, in excitatory responses of muscle, 13, 14 magnesium ions active transport of, 23 and electrically excitable responses in muscle fibre, 23 effect on muscle fibre resting potential, 6 effect on spontaneous miniature pm tentials in muscle, 15, 17 Malacosoma spp., lipid content, 75 malonate in fatty acid synthesis, 132, 133 respiratory inhibition, 122 role in uric acid synthesis, 40,41 Malpighian tubules allantoinase activity, 40 enzyme localization, 58 trehalase activity, 31 I uricase activity, 40 xanthine dehydrogenase activity, 39 maltose, content in locust haemolymph, 292, 295 mammals adipokinetic activity, 184, 185 carbohydrate metabolism, 288, 289, 322 fatty acids, 124, 125, 128, 132, 145147 glycogen metabolism, 330-332, 326 hexokinase, 302 interconversion of non-lipid to lipid, 147 lipids and hormones, 138, 179, 184

SUBJECT INDEX

399

membrane mammals-cont. cellular and subcellular, lipid digestion, 98-100, 113, 115, 184 role of PL, 137, 138, 144 lipid in, 102, 103, 107, 110, 138-140, role of sterols, 176, 178, 180, 209, 143, 144 210 lipid release and transport, 102, 103, cuticular, 152 107, 108, 110, 116, 141, 184 muscle cell, mucopolysaccharides, 341 resting condition, 1 role of sterols, 176 muscle fibre (see Muscle fibre memsugar absorption from gut, 298, 322 brane) Mammestra brassicae, nitrogenous experitrophic, 340, 341 cretion, 55 plasma, 316 manometry, in lipid metabolism studies, postsynaptic (see Postsynaptic mem111-113, 115, 148 brane) manna, trehalose in, 321 membrane permeability mannose muscle fibre absorption and conversion, 298 effect of chemical transmitter, 9 from plasma, glycoprotein, 341 effect of chloride ions, 4, 5, 20 utilization of, 302-304 effect of potassium ions, 4, 5, 11, Mantis, chitin in "silk", 263 effect of sodium ions, 4-6, 11, 22, Mantis religiosa, nitrogenous excretion, 23 46 mass-spectrometry, in isoprenoid stud- membrane potential muscle fibre ies, 168, 181 and inhibitory postsynaptic potenmating, effect of lipids, 86, 169, 186 tials, 19, 20 meconium, composition, 54 and stimulation of excitatory axons, Megoura viciae 8, 10 carbohydrate in haemolymph,291,292 damped oscillation of, 21 trehalase activity in tissues, 311 equation of, 47, 21, 22 Melanoplus bivittatus, nitrogenous extension development, 24 cretion, 46 membrane resistance Melanoplus difserentialis muscle stimulation carbohydrate in haemolymph, 292 effect of blocking agents, 11 eggshell structure, 225,226 effect of carbon dioxide, 6 lipid utilization, 104, 117 effect of GABA, 19,20 sugar absorption from intestine, 298 effect of magnesium ions, 13, 14 trehalase, 310, 311, 313 effect of picrotoxin, 19 trehalose in eggs, 296 Melanoplus sanguinipes, fatty acid con- metabolic energy of uric acid synthesis, 40, 41 tent, 95 metabolic oscillation, in cuticular lamelMelanoplus spp., lipid content, 79 logenesis, 246-254 melezitose, in hemolymph, 295 metabolic pathways, in carbohydrate Meloidae, lipid content, 74 metabolism, 303, 304, 305 Melolontha spp., lipid content, 74 Melolontha spp., trehalase, 311, 314, metabolic switches in cuticular lamellogenesis, 253, 254 315 Melolontha vulgaris, nitrogenous excre- metabolism and resting potential of muscle fibre tion, 50 membranes, 6, 7 Melophagus ovinus, nitrogenous excrecarbohydrate (see Carbohydrate) tion, 52

400

SUBJECT INDEX

metabolism-cont. chitin (see Chitin) glycogen (see Glycogen) lipid, 69-187 (see Lipid for detail) nitrogen (see Nitrogen) trehalose (see Trehalose) metamorphosis and lipids alterations during, 70, 81-89, 96, 142, 143, 146, 172,208 and sterol modification, 174, 175, 180 fatty acid content, 96, 97, 116, 117 glycogen accumulation and conversion during, 327-329,300,301,333, 342, 345,346 mevalonate, as sterol precursor, 161, 165-168, 176 mevalonic acid, in isoprenoid biosynthesis, 161 MGL (see Monoglyceride) Micrelvtra fossularum, nitrogenous excretion, 48 microelectrodes, 10, 12, 15, 19, 20 microfibrils, 214 microorganisms and fat soluble vitamins, 147 and fatty acid synthesis, 133 and sterols, 161, 164, 165, 167, 171, 172, 174, 181 and wax utilization, 101 lipids in, 70 microsomes in lipid metabolism, 98, 137 muscle cell trehalase properties, 314-3 16 microvilli in peritrophic membrane synthesis, 222 mid-gut allantoinase activity, 40 glycogen metabolism, 332, 335 hexokinase activity, 302 mucopolysaccharidein, 341 sugar absorption, 297,298 trehalase activity, 310, 311, 312, 320, 314 trehalose biosynthesis, 306

mid-gut-cont. uricase activity, 40 xanthine dehydrogenase activity, 39 miniature postsynaptic potentials, and release of transmitter substance in muscle, 15-1 7 mitochondria chitin synthetase activity, 344 in lipid metabolism, 89, 118, 119, 121, 122, 124-126, 137-139, 142-144, 166, 167, 176 muscle cell trehalase properties, 314, 315 Moirk effect, 225,228,229 monarch butterfIy, lipid content and behaviour, 86 Monema flavescens glycerol production, 346 glycogen phosphorylase, 333 monoethyloxaloacetate,role in uric acid synthesis, 40 monoglyceride (MGL), in lipid metabolism, 69,97,98, 104, 110, 118, 134 monosaccharides, other than glucose, utilization of, 302-304 (see also Glucose) mormon cricket (see Anabrus simplex) morphogenesis, metabolic oscillators, 246-254 mosquitoes and lipid synthesis, 178, 209 diet and lipids in, 148 motor axons, muscle innervation, 78 moulting and chitin orientation in cuticle, 267, 268 and glucose in haemolymph, 293,323 and glycogen metabolism, 327-329, 341, 342 and trehalase activity, 312, 322, 323 role of chitin metabolism, 328, 341345 role of fatty acids, 97, 102, 117, 145 moulting hormone, and lipids, 171, 172, 177, 179, 180 mucopolysaccharides, 288,328, 340-343 mucosa, intestinal, 98 Mmcu autumnalis, puparium formation, 267

401

S U B J E C T I N D EX

Mmca domestica cuticular lipid, 152, 153 fatty acid content, 95, 96 flight muscle glycogen, 333 trehalase activity, 316 trehalose physiology, 317 glycogen, 327, 333 lipid content, 80 lipid utilization. 110, 111 PL synthesis, 143, 144 sterol modification, 171, 172, 175 uricase localization, 40 muscaterol, 172 Musca vicinia, sterol utilization, 162, 168 muscle and fatty acid synthesis, 132 flight (see Flight muscle) glycogen content, 326 metabolism, 329-331, 333, 334 innervation of, 7, 8, 15,26, 27 insertions in cuticle, 245, 246 leg (see Leg muscle) mammalian glycogen metabolism, 330, 332 skeletal (see Skeletal muscles) thorax (see Thorax muscle) trehalose and trehalase, 307, 310-319, 321 muscle fibre membrane conductance and inhibitory postsynaptic potential, 19 electrical excitability of electrical properties, 20, 21 ionic basis, 21-23 spontaneous activity, 23 mycose, relation to trehalose, 290 Myelobia smerintha, lipid content, 76 Mylabris pustulata, lipid content, 74 myofibrils, trehalase, 315 myosin in muscle contraction, 24, 25 molecular orientation, 214 myristic acid, 92-96, 118, 130 Myzuspersicae, and fatty acid synthesis, 130 Myzus spp., honeydew, 49,92

N NAD (see Nicotinamide adenine dinucleotide) NADP, in lipid metabolism, 69, 119, 128, 129, 133, 164 Nassanoff organ, bee, 169 Nazzara, flight muscles, 14 Neodiprion sertifer, fatty acid content, 95 nerve cord, abdominal, 23 nervous control, chitin orientation in cuticle, 260 nervous system and PL, 138 and sterols, 178 mammal adipokinetic activity, 185 neuromuscular blocking agents, effect on excitory response, 9, 11-13 neuromuscular junctions, transmission process, 17 neuromuscular transmission skeletal excitatory responses, 8-14 inhibitory responses, 17-20 innervation, 7, 8 release of transmitter substances, 15-17 Neuroptera fatty acid content, 94 nitrogenous excretion, 50, 51 neurosecretory cells and carbohydrate metabolism, 338 neurosecretory control over cuticular orientation, 260 Neurospora, trehalase and trehalose, 315, 324 Neurospora crassa, in synthesis of chitin, 343 nicotinamide adenine dinucleotide

WAD)

in ammonia formation, 43 in glutamic acid cycle, 43 in lipid metabolism, 69, 119, 120, 121, 133 in purine oxidation, 37 nitrogen metabolism ,. end products assumptions, 33, 34

402

SUBJECT INDEX

nitrogen metabolism-cont. formation, 34-44 nitrogen content, 34, 35 solubility, 34, 35 nitrogenous excretion amino acids in honeydew, 49 aquatic insects, 47,48, 51, 57 biological significance, 59 Collembola, 45-47 Dermaptera, 46, 48 detoxication of end products, 53, 58 diversity of patterns, 44,45,47 during copulation, 47 during life history, 54, 56 effect of nutrition, 45, 47, 48, 50, 5357

end products, 33-35, 39-44 excretory efficiency, 58 fat body storage excretion, 47, 51 haematophagous insects, 44, 49, 52,

nutrition and blood sugar levels, 291, 292, 295, 299

and lipids conversion from non-lipid, 148,155 fatty acids, 90,92, 93,97,130, 145147

general, 70, 71, 89, 97 isoprenoid compounds, 157-163, 168, 169, 172-176, 179

PL & TGL synthesis, 90, 143, 144 chitin as reserve nutrient, 328, 341 effect on nitrogenous excretion, 45, 47,48, 50, 53-57

utilization of carbohydrate for, 302, 303

Nymphalidae, lipid content, 76 Nymphalis antiopa, fatty acid content, 95 nymphs, lipid content, 73,78,79,81,103 sterol modification, 171, 172

53

Hemiptera, 43,44,48-50 Heteroptera, 48, 49 Homoptera, 49, 50 interpretation, 56-61 Lepidoptera, 54-56 metabolic energy, 53, 58 methods of study, 44, 45 new terminology, 59 Odonata, 4648 Orthoptera, 46, 47 quantitative expression, 45 uricolytic pathway, 35-40 uricotelic pathway, 40, 41 water balance, 57, 58 Noctuidae, lipid content, 75, 76 Nomadacris septemfasciata, chitin orientation, 234 Notodontidae, lipid content, 76 Notonecta glauca, lipid content, 78 nucleic acids in uricolytic pathway, 34-36 oxidative degradation, 35, 42 primary breakdown, 36 nucleosides, in uricolytic pathway, 36, 37

nucleus effect of ecdysone, 180 trehalase, 315

0 Odonata apodeme chitin orientation, 232, 233 daily growth of cuticle, 234 lipid content, 79 nitrogenous excretion, 46-48 odour and biting factor, 160 cockroach, 91 role of terpenes, 160, 169 oenocytes and wax production, 156, 157 PL in, 141 Oestridae, lipid content, 80 oil Cecropia, 177, 180, 183 chrysalis, 168 oleic acid effect on trehalase activity, 314,316 in lipid metabolism, 91-92,94,95-96, 101, 118, 126, 130, 145

Olethreutidae, lipid content, 76 olfaction, and biting, 160 ommochromes, in locust pigmentation, 256

Ornocestus viridulus, chitin orientation, 234

Oncopeltus, lipid content, 78, 117

SUBJECT INDEX

Oncopeltus fasciatus chitin orientation, 234 nitrogenous excretion, 44, 49 pteridine excretion, 44 oocytes, and lipid storage, 185 oogenesis role of hormones, 177, 184, 185 role of lipid, 102, 110, 117, 118, 156, 157, 175 Orconectes viridis, cuticle structure, 227 orientation of chitin-protein complexes, 271 of collagen, 214 of cuticular chitin, 213-279 (see Chitin, orientation) of cuticular protein, 269, 271, 272 of flagellin, 214 of macromolecular polymers, 213, 214 of microfibrils, 214, 215 of myosin, 214 ornithine, precursor of urea, 42 ornithine cycle, 42, 57 Orthocanthacris aegyptium, lipid content, 78 Orthoptera circadian clock, 234 cuticle structure, 226, 234 fatty acid synthesis, 131 lipid content, 78, 79, 95, 142 nitrogenous excretion, 46,47 sterol utilization, 160 uricolytic enzymes, 37, 46, 47 Oryctes, 18 Oryctes nasicornis, lipid content, 74 Oryzaephilus, ornithine cycle, 42 oscillation, in morphogenesis, 246-254 ovary and action of juvenile hormone, 184, 185 and glycogen metabolism, 335, 339, 340 lipids content, 117 uptake of sterol, 172, 173, 174 overwintering, role of carbohydrates, 296,340,345 ovipositors, parallel chitin, 220 oxalate, and trehalase activity, 322 Oxya japonica, lipid content, 78

403

P 32Porthophosphorus, in PL studies, 142, 144 Pachymeris, lipid content, 73,89 [14C]palmitate, in carbohydrate metabolism studies, 337 palmitic acid, in lipid metabolism, 91, 94-96,101, 126,128,130,132 palmitoleic acid, in lipid metabolism, 92,94,95, 129,130 Palomena prasina, nitrogenous excretion, 48 pancreas adipokinetic activity, 185 lipase activity, 113,115, 116 Papilio spp., lipid content, 76 parabolic lamellae, in cuticular chitin, 223-229 passive diffusion (see Diffusion) Pectinophora gossypiella, lipid content, 75 Pegomyia ulmaria, lipid content, 79 Pemphigus populicaulis, fatty acid content, 94 Pemphigus utricularias, lipid content, 78 peripheral inhibition, in muscle nervous response, 17, 18 Periplaneta glutamic dehydrogeriase, 43 leg muscle glutamate and contraction, 12 ion basis of electrically excitable responses, 22 membrane potential, 2, 4 muscle postsynaptic potentials, 13, 15 resting potential, 5,6 sterols, 171, 172 xanthine dehydrogenase, 39 Periplaneta americana amino acids in cuticle, 275 calcium in ootheca, 276 carbohydrate metabolism effect of hormones, 338,339 glycogen, 326, 331 glycoproteins, 341, 343 haemolymph, 292 sugar absorption, 297 chitinase activity, 345

404

SUBJECT I N D E X

PerQlanef a americana-cont. chitin orientation, 234 circadian clock, 234, 239 flight muscle, 316, 318 lipids content, 79, 118, 208, 209 cuticular, 152-155 effect of hormones, 185 fatty acids, 97, 209 utilization, 99-101, 103, 104 nitrogenous excretion, 46 photosensitivity, 257 uric acid biosynthesis, 40, 41 peritrophic membrane crossed fibrillar structure, 222 extracellular polymerization, 263 glycoproteins, 340, 341 synthesis, 222, 263 Pezotettix giornae, lipid content, 79 PH and fatty acid oxidation, 122, 123, 125 effect on lipid hydrolysis, 112, 114 phagostimulants, 160 Phalera bucephala fatty acid content, 96 lipid content, 76 pharmacology, and excitation of skeletal muscle, 15, 16, 19,20 Phasmida, chitin orientation, 234 pheromones, 70, 169, 177, 186 Philaenus spumarius, fatty acid content, 94 Phormia spp. carbohydrate metabolism and flight, 297, 311, 316-318 effect of hormones, 338 glucose, 301 glycogen, 326 haemolymph, 291, 294, 322, 323 sugar absorption and levels, 297299 trehalase, 311, 313, 315, 316, 322, 323 trehalose, 304, 306, 307, 317, 318, 321 use of monosaccharides, 303 lipid content, 80, 99 Phormia regina PLin, 143

Phormia regina-cont. sterol utilization, 162 phosphatases in sugar biosynthesis and utilization, 301, 302, 305, 307 phosphatidylcholine (PTC), in lipid metabolism, 138, 140-144,209 phosphatidylethanolamine (PTE), in lipid metabolism, 69, 138, 140-144, 209 phospholipid (PL) synthesis general mechanism, 132-137 metabolism and function, 137-144, 160, 180, 185,209 phosphorylase, glycogen, 305, 326, 329, 331-334,337,339 photoperiod (see Light) photoperiodic response, 255 photosensitivity, and chitin orientation, 254-257 phragmata daily growth layers, 245,246 Phyllophaga rugosa, lipid content, 74 phytol, juvenile hormone activity of, 180, 181 phytosterols, 173, 174, 178 picrotoxin, effect on inhibitory responses in muscle, 19 Pieris brassicae, nitrogenous excretion, 55 Pieris spp., lipid content, 76 pigmentation, cuticle, 256, 257 Pissodes, flight muscle, 6 Pissodes notatus, trehalase activity, 311 Pituitary gland, adipokinetic activity, 185 PL (see Phospholipid) Plagionotus arcuatus, lipid content, 73 plasma, glycoproteins, 341, 343 plasma membrane, and trehalase location, 316 Plumatella, trehalose in, 325 Plusia gamma, lipid content, 76 Podura aquatica cuticle structure, 227 epidermal brush border, 278 extracellular cuticle formation, 264 265 poisons, metabolic, 298

SUBJECT I N D E X

polarization analysis in age determination, 245, 246 of chitin orientation, 217, 222-225, 247,258 of locust cuticle, 235, 236, 247, 258 pollen, sterol in, 168 polyamides, microfibril diameter, 214 polyethylene plastic, microfibril diameter, 214 poIymer linkages, tensiIe strength, 217 polymerization extracellular, 263,264 in chitin synthesis, 261,263,264 of cellulose, 263 polyols, 296 polysaccharides biochemistry of, 301-336 macromolecular comparison, 213214 Popillia japonica lipid content, 74 uricase, 38 population, age dynamics, 245 pore canals cuticular arrangement, 226 in reoriented cuticle, 268, 269, 270 postsynaptic membrane muscle and quantal release transmitter substance, 17 magnesium reduction in sensitivity, 13, 15, 17 permeability in muscle action of inhibitory transmitter substance, 19,20 action of GABA, 19, 20 to chloride ions, 20 postsynaptic potential in muscle excitatory response and calcium-magnesium antagonism, 13, 14 and stimulation of “fast” and “slow” axons, 8-1 1 effect of blocking agents, 11, 12 ionic basis of, 9-1 1 time-dependent properties of, 14 muscle and inhibitory response, 17-20

405

postsynaptic potential-conr. as stimulus for electrically excited response, 20,21 effect of transmitter substance, 9, 10, 13-17 quantal nature of, 13,14,17 spontaneous miniature, 15-17 potassium conductance, in muscle fibre membrane, 21,22 potassium contracture, comparison of “fast”and “slow”muscle fibres, 26,27 potassium electrode hypothesis, muscle cell membrane, 4 potassium ion and depolarization and contraction of muscle, 24-27 effect on inhibitory postsynaptic potentials in muscle, 19 effect on muscle fibre membrane permeability, 4,5, 11 effect on muscle resting potential, 2-4, 7 effect on spontaneous miniature potentials in muscle, 15 potato beetle, trehalase in, 310 potential, resting (see Resting potential) potentials, postsynaptic (see Postsynaptic potentials) prealar arm, rubber-like cuticle, 2152 16,246-249 precursors for vitamin D, 172 in chitin metabolism, 343, 344 in lipid metabolism, 145, 148, 159, 161,171,176,179,209 of urea, 42 of uric acid, 40, 41 pregnenolone, 173 prepupa, lipid content, 74,75,77,80-83, 96.97 presynaptic nerve ending, and release of transmitter substances, 15, 17 presynaptic potential muscle and calcium-magnesium .antagonism, 13 and action of Wbon dioxide, 14 and frequency of miniature postsynaptic potentials, 15

406

SUBJECT INDEX

Procrustes coriaceus, nitrogenous excretion, 50 Prodenia, and lipid metabolism, 150, 209 Prodenia eridanica chitin synthesis, 344 fatty acid synthesis, 131, 132 glyoxylate cycle enzyme, 329 Prodenia ornithogalli, lipid content, 75 progesterone, 173 propionate, effect on inhibitory postsynaptic potentials in muscle, 19 prostigmine, effect of spontaneousminiature potentials in muscle, 16 proteins chitin metabolism, 341-345 deamination, 42,43 glycoproteins, 340, 341 in ammonia metabolism, 43 in lipid metabolism, 85, 87, 102, 103, 108-110, 114, 116, 127, 128, 137, 139,140,148,176,178,209 intake and uric acid deposition, 47 proteins orientation in cuticle, 269, 271, 272 oxidative degradation, 34,35,40 prothoracic gland and sterols, 172, 178 effect on carbohydrate metabolism, 336,338 Pseudococcus citri, uric acid in honeydew, 49 Pseudophonus pubescens, lipid content, 73 Psychidae, lipid content, 76 PTC (see Phosphatidylcholine) PTE (seePhosphatidylethanolamine) pteridines, excretion, 44 Pteronidea spp. nitrogenous excretion, 52 Pterostichus spp., lipid content, 73 Ptinus tectus, sterol utilization, 162 pupa, diapause (see Diapause) puparium cuticular structure, 223,267 purine biosynthesis, 37,40,41 deamination, 34,42 oxidation, 34,37

purine nucleotidase, 36,37 Pyralidae, lipid content, 76,77 Pyrausta nubilalis lipids, 77, 163 sterol utilization, 163 pyrimidines, metabolism, 35, 36 Pyropyga decipiens, fatty acid content, 94 Pyrrhocoris apterus carbohydrate metabolism, 338 lipid content, 78, 89 [14C]pyruvate in trehalose synthesis studies, 305

Q

Qlo of endocuticular growth, 244, 245 Qlo of circadian clock, 239, 244, 245 quanta1release, of transmitter substance in neuromuscular transmission, 13-17 queen bee, larvae, lipids in, 92, 93, 168 queen substance, 186

R radioisotopes tritiated tyrosine, 234 use in studies biosynthesis of trehalose, 304-307 chitin metabolism, 342-344 fatty acids, 117, 121, 123, 124, 130133 glycogen metabolism, 330, 331 hydrocarbons and waxes, 155, 156 isoprenoid compounds, 161, 167, 168, 171, 172, 179-181, 185 lipid metabolism, 104,131,148,150 lipid release and transport, 103, 104, 106, 108 nitrogen excretion, 53 phospholipid and triglyceride, 142, 144 substrate interconversion, 147-151 sugar absorption and regulation, 297-300 uric acid synthesis, 40,57 ramie, tensile strength, 219 Rana, muscle resting potential, 6 rat liver, lipid metabolism and hormones, 184 and mitochondria, 139, 176

SUBJECT INDEX

rayon, microfibril diameter, 214 reducing substances, and glucose occurrence, 289 refractoriness, in excitation of skeletal muscle, 14, 21 regeneration, and PL, 138 “relaxing factor”, in muscle contraction, 25

release lipid, 102-111, 117, 119, 209 of transmitter substance in neuromuscular transmission, 15-17 reproduction, effect of lipids in diet, 145, 209

reptiles, ornithine cycle, 42 resilin chemical and mechanical properties, 217

content of rubber-like cuticle, 216, 248

daily growth layers, 234, 246-249, 252. 253

in cuticle,216,234,246-249,252-254, 272,273

macromolecular network, 272-273 optical density, 273 respiration effect of hormones, 337, 340 flight muscle, 118-127 respiratory quotient and carbohydrate conversion to lipid, 148

and fight, 119, 121 and sexual dimorphism in lipid metabolism, 87, 88 responses excitatory (see Excitatory responses) inhibitory (see Inhibitory responses) resting potential muscle membrane effect of carbon dioxide, 6 effect of temperature, 6, 7 effect of various ions, 2,4-7 reticulum endoplasmic and “relaxing factor”, 25 and trehalase, 316 sarcoplasmic and trehalase, 315, 316

407

reversal potential, 19 Rhagium inquisitor, lipid content, 73 Rhaphigaster griseus, nitrogenous excretion, 48 Rhinocricus nodul@es, cuticle structure, 227 Rhodnius creatin excretion, 44 cuticle expansion, 263 cuticle hydration, 277, 278 endocuticle and nutrition, 341 epicuticle expansion, 268 intermoult tracheal growth, 268 Rhodnius prolixus nervous control over growth, 260 nitrogenous excretion, 48 Rhoecocoris sulieventris, scent, 155 Rhynchophorus palmarum, lipid content, 74 ribose-5-phosphate, in purine synthesis, 4041

rice stem borer, sterol synthesis, 161 Romalea, muscle, effect of ions on potentials, 5, 6 inhibitory responses, 18-20 Rothschildia spp., lipid content, 77 royal jelly, lipid in, 92, 93, 168 S saliva, chitinase activity, 345 salivary gland glucose-6-phosphatase,301 sugar levels, 296 trehalase activity, 310, 311, 319 Samia Cynthia

carbohydrate in haemolymph, 293 glycogen metabolism, 327, 333 lipids in, 77, 154, 155, 180 Samia Cynthia ricini

juvenile hormone extracts, 183 trehalase activity in tissues, 312, 320, 323 Sarcophaga

exocuticle lamellogenesis, 264 flight muscle, 6, 316 sarcosomes, flight muscle (see Mitochondria). ,. Saturnia pyri, lipid content, 77 Saturniidae, and lipids, 77, 178

408

SUBJECT I N D E X

scale production, and fatty acids in diet, 145 scales, chitin orientation in cuticle, 222, 263 Scarabaeidae chitin orientation, 221 lipid content, 74 scent secretion, lipids in, 155, 169 Schistocerca carbohydrate metabolism chitin synthesis, 343 fat body, 330 haemolymph, 292 monosaccharide utilization, 303 sugar absorption, 297, 298 trehalose biosynthesis, 304,306,307 flight muscle innervation, 8 leg muscle and calcium-magnesium antagonism, 13 blocking effects of compounds on excitatory responses, 8, 9, 10, 11, 12 inhibitory responses, 18-20 innervation, 8 ions and contraction, 24, 25 potassium ions and membrane potential, 3, 4 spontaneous miniature postsynaptic potentials, 15 spiracular muscle and excitatory response, 8,9 effect of carbon dioxide, 6, 14,26 Schistocerca gregaria chitin orientation, 234-260 circadian clock, 234, 239 endocuticle structure, 236-238 exocuticle structure, 235-236 experimental use of cuticle, 235 lipids and flight, 116, 119 content, 79 fatty acids in diet, 145 sterols, 160-162 nitrogenous excretion, 46 Qlo of lamellogenesis, 239, 244, 245 uncoupling lamellogenesis, 241 Schistocerca paranamense, chitin orientation, 234

Schistocerca spp., implantation of dermal fragments, 259 Schmidt’s layer, 265 Sciara coprophila, glucose-6-phosphatase, 301 Scoliopteryx libatrix, lipid content, 76 secretion defensive, 169, 210 lipid in 155, 169, 209 Selaginella, trehalose in, 291 Semiadalia undecimnotata, glycogen in over wintering adults, 346 sex, and lipase activity, 112, 115 sex attractants, 70, 180, 186 sex glands, accessory, 47 sexual dimorphism and lipid metabolism ant odour, 169 content, 81, 84-89, 96, 97, 99 esterase activity, 115 extra-cuticular hydrocarbon, 156 sexual maturation, role of monoterpenoids, 160, 161 sialic acid, 341 Sialis lutaria, nitrogenous excretion, 51 silk gland and carbohydrate metabolism, 312. 321, 323

PL in, 142 silkworm, adenase and guanase, 37, 41 and sterol biosynthesis, 167, 168, 174 biting factor, 160 glycogen level during starvation, 299 hexokinase activity, 302 sugar in haemolymph, 291 trehalose, 299, 305 uricotelic pathway, 41 Siluanus surinamensis, sterols utilization 162 silver fish (see Ctenolepisma) simple lipids, 72 Siphyloidea sp., chitin orientation, 234 Sitona scissifrons, fatty acid content, 94 sitosterol, in lipid metabolism as growth factor, 162, 163 brain hormone, 177, 178 content, 168 sterol modification, 171-174 structure, 158, 160

SUBJECT INDEX

size, and lipid content, 81-85 skeletal muscles, excitation of, 1-27 excitability of muscle fibre membrane, 20-23

excitation-contraction coupling proC ~ S S , 23-27

neuromuscular transmission, 7-20 resting potential, 2-7 skeleton, differentiation, 213 skin, and fatty acids in diet, 145 slime mould trehalase, 315 trehalose in, 324 “slow” and “fast” axons (see “Fast” and “slow” axons) snail muscle, glutamate and contraction, snake venom, effect on trehalase activity, 316

sodium, conductance in muscle fibre membrane, 21, 22 sodium extrusion mechanism, 6, 7 sodium ions and permeability of muscle fibre membrane 4-6, 11, 22, 23 effect on muscle resting potential 2,4, 5,6, 7

sodium ion transport, and PL, 138, 180 sodium pump, 7, 180 sorbitol production, 325,345, 346 sorbose, utilization, 303 spectrometry, infra-red, in cuticular wax studies, 153 spectrophotometry in demonstrating glycogen synthetase, 331

in lipase study, 111 spectroscopy,infra-red in lipid studies, 186, 187 of chitin orientation, 217 spermathecae, and uric acid, 47 spermatophore, chitin in, 263,264 Sphenarium purpurescens, lipid content, 79 Sphinx ligustri, Carbohydrate in hemolymph, 293 Sphinx spp., lipid content, 77, 78, 209 spiracular muscle effect of carbon dioxide on 6,14,26

409

spiracular muscle-cont. excitatory response, 8, 9 potassium ions and contraction, 24 spontaneous activity of membrane, 23 Spodoptera abyssinia, lipid content, 76 spontaneous activity, and excitability of muscle fibre membrane, 23 spontaneous miniature postsynaptic potentials, 15-17 squalene, in sterol biosynthesis, 161, 165, 166 squid, giant axon, 5 starvation and cuticle reabsorption, 341, 345 and ‘lipid metabolism, 89, 100, 141, 142 effect on carbohydrate levels, 299, 306, 324, 329 stearic acid, in lipid metabolism, 91, 92, 94-96, 129, 130 Stegobium peniceum, sterol utilization, 162 Stenobothrus lineatus, chitin orientation, 234 Sternocera castanea, elytra structure, 232 sterols as growth factors, 162, 163 biosynthesis, 161,164-167,176,209 content, 104,105,118,168,169 function 103, 147, 175, 176, 178, 180, 209,210 in cuticular wax, 153, 154 in nutrition, 157, 159, 160, 209 modification, 170-175 structure, 158, 159 Stictocephala diceros, fatty acid content, 94 stigmasterol, as growth factor, 162, 163 content in insects, 168 in hormones, 177 in sterol modification, 174, 175 storage, lipid, 99, 102-106,110,146,148, 172,178,185,209,210 storage excretion, uric acid, 47, 51 Streptomyces antibioticus, chitinase production, 344, 345 strontium, as substitute for calcium in musclewontraction, 25, 26

410

SUBJECT I N D E X

sub-oesophageal ganglion, dispausehormone, 339,340 substrate interconversion, in lipid biosynthesis, 147-152 sucrase, pH in mid-gut, 320 sucrose effect on muscle resting potential, 5 effect on spontaneous miniature potentials in muscle, 15 in fatty acid synthesis, 130 in haemolymph, 291,292,295 in whole insects, 296 utilization of, 303 sugar content and developmental stage, 291-297, 309, 323, 324 and frost-resistance, 296 of haemolymph, 291-295 of insect tissues, 296, 297 of whole insects, 295, 296 sugars active transport of, 298, 322 biochemistry of, 287-347 biosynthesis and utilization 298, 301325 intestinal absorption, 297-299, 320 occurrence, 289-297 regulation in blood, 299-301,309, 329 survival and fatty acids, 145 and glycerol and sorbitol production, 325, 346 and monosaccharide utilization, 302, 303 and sugar levels, 296, 299 symbionts and fatty acid synthesis, 130, 131, 160, 167 swimming hairs, parallel chitin, 220 and trehalase activity, 320 in sterol biosynthesis, 160, 167, 168 synapticdelay, effect of tryptamine, 12 synthetase, chitin, 343, 344 glycogen, 305, 330-331 trehalose, 307-309,337 synthetic polymers, microfibril formation, 214 Syrphidae, lipid content, 80 systematics, and lipid content, 89

T Tanytarsus lewisi, lipid content, 80 techniques autoradiography (see Autoradiogwhy) chromatography (see Chromatography) electron microscopy, chitin orientation in cuticle studies, 223, 229, 236, 239, 246, 264, 265 (see also Electron microscopy) electrophoresis(see Electrophoresis) extraction, glycogen, 326 lipid, 71-73 for carbohydrate analysis, anthrone reaction, 289, 294, 300, 323 chromatography,289,294-296,304, 307 enzymatic assays, 289, 294, 296, 307, 332 histochemistry (see Histochemistry) implantation experiments, in cuticle lamellogenesis studies, 257-260 in blood sugar regulation studies, 299, 300 in chitin metabolism studies, 342-345 infra-red spectrometry, cuticular wax studies, 153 in lipid studies, 186, 187 of chitin orientation, 217 in glycogen phosphorylase activity studies, 333, 334 in glycogen synthetase demonstration, 330, 331 in insect biochemistry, general discussion, 288, 289 in sugar absorption studies, 297, 298 in trehalase studies, 312,313,315,316 lipase assay, 111, 112, 113, 115 manometry, in lipid metabolism studies, 111113, 115, 148 mass spectrometry, in isoprenoid studies, 168, 181 purification, of hormones, 177, 180

SUBJECT INDEX

techniques-cont. xanthine dehydrogenase, 56 radioisotopes (see Radioisotopes) spectrophotometry, in demonstrating glycogen synthetase, 331 in lipase study, 111 staining, chitosan-iodide and exocuticle, 235, 238 testing applicability of resting potential equation, 2, 3, 4 treatment with EDTA of muscle contraction, 25 use of microelectrodes in muscle, 10, 12, 15, 19, 20 X-ray diffraction, in chitin studies, 217, 223, 266, 267, 27 1 Telea, spiracular muscles, 6 temperature and excitation of muscle, 6, 7, 9 and lipid metabolism, 89, 93, 96, 209 effects on chitin orientation, 239, 241, 242,244,245,255 temperature coefficient, circadian clock, 239,244,245 tendons, parallel chitin, 220 Tenebrio juvenile hormone extract, 180 lipids, content, 74 fatty acids, 145 metabolism, 117, 163, 209 muscle membrane effect of carbon dioxide, 6 potentials, 4 xanthine dehydrogenase, 39 Tenebrio molitor carbohydrate in haemolymph, 294 cuticle structure, 226 glycogen and development, 327 nitrogen metabolism, 51 Tenthredinidae, lipid content, 81 termite chitinase activity, 345 isoprenoid content, 169 terpenes and juvenile hormone, 180

411

terpenes-cont. in insects, 169, 176, 209 in nutrition, 157, 160, 209 tetanic tension, 20 Tetraopes tetraophthalmus, fatty acid content, 94 Tettigonia viridissima, chitin orientation, 234 lipid content, 79 TGL (see Triglyceride) Thaumetopoeapityocampa, lipid content, 78 thawing and freezing, effect on trehalase activity, 314, 316, 319 Thermobia domestica, and sterol biosynthesis, 167 thorax ligation effect on carbohydrate metabolism, 338 muscle flight and carbohydrate levels, 318, 329 glycogen phosphorylase activity, 334 properties of trehalase, 314, 316 Thyridopteryx sp., lipid content, 76 tibia, cuticular structure, 240 time-dependent properties of postsynaptic potentials in muscle, 14 Tineola bisselliella, nitrogenous excretion, 56 Tipula paludosa, nitrogen metabolism, 52 tissues amylase activity, 334, 335 chitinolytic activity, 345 glycogen phosphorylase activity, 332334 glycogen synthetase activity, 330, 331 interrelation of in carbohydrate metabolism, 321 sugar levels, 296, 297 trehalase activity, 310-312, 316, 318, 319, 321, 322, 328 urea in, 41 trachael taenidia, chitin orientation, 222 transamination, in ammonia formation, 43

412

SUBJECT INDEX

transmission,neuromuscular(see Neuromuscular transmission) transmitter substance acetylcholine effect on postsynaptic potential in muscle, 9, 10, 15 effect on excitatory responses of muscle nature of, 11-14 inhibitory and GABA, 19,20 quantal release in neuromuscular transmission, 15-1 7 quantal nature of postsynaptic potential, 17 spontaneousminiature postsynaptic potentials, 15, 17 transpiration, role of cuticular lipid, 152-154 transport active of sugars, 298, 322 lipid, 97-99, 102-110, 116, 126, 127, 141, 184 trehala, 290 trehalase activity in tissues, 310-312, 316, 318, 319, 321, 322, 328 and moulting, 312, 322, 323 and trehalose physiology (see Trehalose) characteristics of, 313-317 in trehalose cleavage and use, 309-3 16 non-insect, 315 precise location of, 315, 316 trehalose and chitin metabolism, 261, 343 and trehalase physiology haemolymph, 322-324, 321 intestine, 319-322 muscle, 317-319, 321, 329 biosynthesis of enzymes of, 305-309 synthesis of tissues and homogenates, 306, 307 synthesis in vivo, 298, 304-306, 321 characteristics of, 289, 290, 325 cleavage and use of, 309-316, 321 dormancy and properties of, 324,325

trehalose-cont. hydrolysis, 305, 309-31 6, 318, 321 levels effect of hormones, 336-340 in haemolymph, 292-297 in tissues, 296, 297 in whole insects, 295, 296 regulation in blood, 297-301, 309, 329, 337 occurrence, 289-291, 324 triacetin, hydrolysis of, 111, 112, 115 Tribolium confusum, and lipid metabolism, 117, 162, 173 tributyrin, hydrolysis of, 111, 112, 115, 116 Trichiocampus populi, sugar levels and overwintering, 296 Trichoplusia ni, fatty acids in diet, 146 Trichoptera, fatty acid content, 94 triglyceride (TGL) content in insects, 89,90,92, 118, 178 digestion and absorption, 97-99, 102, 111, 113, 116 metabolism and function, 118, 139144, 175, 178, 184, 185,209 release and transport, 103-108, 110, 184 synthesis, 133-137, 148 Trigonophora meticulosa, nitrogenous excretion, 55 triolein, hydrolysis, 100, 101, 113-1 15 Trirhabda virgata, fatty acid content, 94 trisaccharides,absorption from gut, 299, 320 tritium incorporation into chitin, 265 incorporation into resilin, 234 trityrosine, in resilin orientation, 234 tryptamine, blocking effect of on excitatory responses of muscle, 11, 12, 21 tsetse fly, haematin excretion, 44 (see also Glossina) TSH, sensitivity of lipase, 184 Tuberolachnussalignum fatty acid content, 94 nitrogenous excretion, 49 tubocurarine, effect on muscle excitatory response, 12

SUBJECT INDEX

tunicates cellulose microfibrils, 214 macromolecular orientation, 214, 223 microfibril orientation, 223-225, 227, 229 tunicin comparison with cellulose and chitin, 213, 214,227 parabolic lamellar structure, 223 twitch tension, in muscle, 9, 20, 24, 26

413

uric acid-cont. Odonta, 4648 Orthoptera, 46,47 fat body deposits, 47, 51 in accessory sex glands, 47 in uricolytic pathway, 35, 36, 38 in uricotelic pathway, 40,41 nitrogen reserve, 47 precursors of, 40,41 uricase, effect on, 37, 38 uricase Collembola, 45 U distribution in insects, 37, 38, 47 U D P (see Uridine diphosphate) Hemiptera, 48 ultra-violet chromatography, in lipid in uricolytic pathway, 37-40,45 studies, 172 localization in insects, 3940 uncouplers, and glyceride synthesis, 108 uricolytic enzymes urea adenase, 36,37 biosynthesis, 39,41,42,49 adenosine deaminase, 35, 37 enzymic decomposition, 39 allantoicase, 38, 39, 45 excretion allantoinase, 3840,45 in various orders, 46, 48, 49, 52, 53 during life history, 54 pathways, 34,35, 38,41 guanase, 36,37 role, 56, 57 guanosine deaminase, 36, 37 origin, 57 in Coleoptera, 50, 51 role in uric acid synthesis, 40, 41 in Collembola, 45 soluble end product, 34 in Hemiptera, 48 urease in Homoptera, 50 in uricolytic pathway, 38, 39 in Hymenoptera, 52 ureotelic insects, excretory terminology, in Lepidoptera, 56 59 uric acid in Orthoptera, 37, 46, 47 methods of study, 56 and copulation, 47 nonexcretory €unctions, 59 biosynthesis, 36, 37,40, 41,57 purine nucleotidase, 36 degradation, 37-39 urease, 38, 39 end product uricase, 3740,45 protein metabolism, 30, 40, 53, 57, xanthine dehydrogenase,36,37,39,56 58 xanthine oxidase, 37 purine metabolism, 48 uricolytic pathway excretion components, 35-36,38 Coleoptera, 50, 51 definition, 34 Dermaptera, 46,48 discussion, 3640 Diptera, 52-54 enzymes (see Uricolytic enzymes) Hemiptera, 48-50 nucleoside deamination, 36 Heteroptera, 48, 49 uricotelic detoxication, metabolic cost, Homoptera, 49, 50 53,58 Hymenoptera, 51,52 uricotelic pathway, components, 35 Lepidoptera, 54-56 uricotelic pathway mechanisms, 33,34,40 definition, 34 Neuroptera, 50,51

414

S U B J E C T INDEX

uricotelic pathway-cont. discussion, 40-41 enzymes, 40,41 uricotelism adaptation to terrestrial life, 33 definition, 60 development in insects, 59 generalization, 57 uridine diphosphate (UDP), in chitin synthesis, 261 uridine triphosphate (UTP),in chitin synthesis, 261 Uroleon nostras, introgenous excretion, 50,51 uronic acid, 341 utilization glucose, 301, 302 monosaccharides, other than glucose, 302-304 trehalose, 309-316 UTP (see Uridine triphosphate) utriculi majores, uric acid storage, 47

V

van der Waal's forces in chitin structure, 217, 218 Vanessa atalanta, nitrogenous excretion, 55 Vanessa urticae, lipid content, 76 venom, snake, effect on trehalase activity, 316 ventriculus, sugar absorption, 297 Verlusia rhombea, nitrogenous excretion, 48 vertebrate hormones effect on lipid metabolism, 184-186 in insects, 210 vertebrate muscle contraction, 23-25 miniature postsynaptic potentials, 15 resting potential, 6 vertebrates blood sugar levels, 291 glycogen phosphorylases, 332, 333 macromolecular orientation, 214 Vespa cincta, lipid content, 81 Vespuk, flight muscle, 6

vision, and carotenoid synthesis, 164 vitamin D, precursor, 172

W walking stick, defensive secretion, 209 water active transport in gut, 298 and lipid content, 85, 86, 116 effect on chitin tensile strength, 218 water absorption and chitin orientation in cuticle, 277 water beetle, cortisone in, 210 water loss, role of waxes and hydrocarbons, 152-154, 156 water shortage, effect on excretion, 57 WaX

composition of, 93 cuticular, 93, 152-155 effect of hormones on production, 155 extra-cuticular, 155-1 57 in diet, 100, 101, 140, 141 in lipid classification, 72 wing, chitin synthesis in, 343, 344 wing beat frequency, and trehalose levels, 317-3 19 wing hinge ligament, cuticular structure, 253,254 wing veins, parallel chitin, 220

X xanthine enzymic oxidation, 37 excretion, 52, 56 in protein metabolism, 41, 58 in uricolytic pathway, 35, 36 in uricotelic pathway, 41 xanthine dehydrogenase, 36,37,39,56 xanthine oxidase, 37, 41 Xenylla welchii, allantoicase activity, 39 X-ray diffraction, in chitin studies, 217, 223,266,267,271 xylose from plasma glycoprotein, 341 utilization, 303

Y

Yeast, and fatty acid synthesis, 129

415

SUBJECT INDEX

yeast trehalases, 3 15 trehalose metabolism, 290 Young's modulus, of cuticle, 215

Z

zymosterol in cholesterol biosynthesis, 165 structure, 159

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