neurohormonal techniques in insects

Thomas A. Miller. Editor
Neurohormonal Techniques in Insects
With a Foreword by Gottfried S. Fraenkel
With Contributions by R. J. Aston . T. Goto . L. Hughes· H. Ishizaki
M. Isobe . K. J. Kramer' S. H. P. Maddrell W. Mordue . S. E. Reynolds· I. M. Seligman
A. N. Starratt . R. W. Steele· J. V. Stone· A. Suzuki J. W. Truman' J. zditrek
Springer-Verlag [$] New York Heidelberg Berlin
Thomas A. Miller Department of Entomology University of California Riverside, California 92521
With 90 Figures
Library of Congress Cataloging in Publication Data Main entry under title: Neurohormonal techniques in insects
(Springer series in experimental entomology) Bibliography: p. Includes index. I. Insect hormones. 2. Neurosecretion. I. Miller,
Thomas A. II. Series. QL495.N48 595.7'01'88 79-27343
All rights reserved. No part of this book may be translated or reproduced in any form without written permission from Springer-Verlag. The use of general descriptive names, trade names, trademarks, etc. in this publication, even if the former are not especially identified, is not to be taken as a sign that such names, as understood by the Trade Marks and Merchandise Marks Act, may accordingly be used freely by anyone.
© 1980 by Springer-Verlag New York Inc.
Softcover reprint of the hardcover 18t edition 1980
987654321
e-ISBN-13: 978-1-4612-6039-4
Series Preface
Insects as a group occupy a middle ground in the biosphere between bac teria and viruses at one extreme, amphibians and mammals at the other. The size and general nature of insects present special problems to the student of entomology. For example, many commercially available in struments are geared to measure in grams, while the forces commonly en countered in studying insects are in the milligram range. Therefore, tech niques developed in the study of insects or in those fields concerned with the control of insect pests are often unique.
Methods for measuring things are common to all sciences. Advances sometimes depend more on how something was done than on what was measured; indeed a given field often progresses from one technique to another as new methods are discovered, developed, and modified. Just as often, some of these techniques find their way into the classroom when the problems involved have been sufficiently ironed out to permit students to master the manipulations in a few laboratory periods.
Many specialized techniques are confined to one specific research labo ratory. Although methods may be considered commonplace where they are used, in another context even the simplest procedures may save con siderable time. It is the purpose of this series (1) to report new develop ments in methodology, (2) to reveal sources of groups who have dealt with and solved particular entomological problems, and (3) to describe ex periments which might be applicable for use in biology laboratory courses.
THOMAS A. MILLER, Series Editor
Call to Authors
Springer Series in Experimental Entomology will be published in future volumes as contributed chapters. Subjects will be gathered in specific areas to keep volumes cohesive.
Correspondence concerning contributions to the series should be com municated to:
Thomas A. MiIIer, Editor Springer Series in Experimental Entomology Department of Entomology University of California Riverside, California 92521 USA
Foreword and Overview
It should be emphasized from the outset what this book is meant and what it is not meant to be. It brings together the very considerable and diffuse information about neurohormones in insects largely from the point of view of the hard facts-evidence for their existence, their chemical na ture, and the techniques used in obtaining this information. In this re spect, it is invaluable to everyone entering this field and despairing how to pick the right insect and method out of a seemingly infinite variety of choices. The book does not give an integrated picture ofthe interaction of these hormones, and omits to tell the often strange and exciting stories of the devious ways by which these hormones were discovered.
What gives this volume a certain distinction and authority, different from similar ventures, is the fact that most chapters were written by the very person or group that made the original discoveries, worked out the original methods, and are still active in the field.
Classification of Insect Neurohormones
This book deals with an almost bewildering variety of neurohormonal manifestations, which makes the reader wonder about how to view them in an orderly scheme. A classification has recently been devised by Seh naP and is given here in somewhat abbreviated form (translated from the German):
1 Sehnal F. (1979). Neuroendokrine. Regulation der Entwicklung der Lepidop teren. In, Probleme der Korrelation neuraler und endokriner Regulation bei Ever tebraten. Ed. H. Penzlin. Wissenschaftliche Beitrage der Friedrich-Schiller Universitat, Jena, 154-175.
X Foreword and Overview
a) Glandotropic neurohormones guide the activity of endocrine glands, viz. prothoracicotropic hormone (Chap. II), allatotropic hormone.
b) Morphogenetic neurohormones guide the speed and direction of ontogenesis, i.e., shape, structure, color, viz. bursicon (Chap. 5), pupariation hormones (Chap. 7), diapause hormone (Chap. II).
c) Myotropic neurohormones affect the kinetics of the heart, intes tine, the Malpighian tubules, the oviducts, ovaries and other inter nal organs, viz. proctolin (Chap. I).
d) Metabolic neurohormones influence metabolism, viz. adipokinet ic hormone (Chap. 2), insulin-like hormones (Chap. 5), diuretic hormone (Chaps. 3 and 4).
e) C hromotropic hormones affect rapid color change by migration of pigment (rather rare in insects, not dealt with in this book).
f) Ethotopic neurohormones act on the nervous system, viz. eclosion hormone (Chap. 9), the pupariation factors (Chap. 7).
Historical Background
Unlike vertebrate endocrinology, which has developed largely through the study of the control of individual growth and metabolic processes, in sect endocrinology developed almost exclusively from the study of complicated morphogenetic events, such as molting and metamorphosis. The latter turned out to be controlled largely by the two master glands, the corpus allatum and the prothoracic gland, with an overall control by neurohormones. In this sense, vertebrate endocrinology was from the beginning biochemically oriented, while insect endocrinology largely stemmed from a study of morphology and developmental physiology.
This preoccupation with the hormonal control of developmental events so dominated insect endocrinology, including neuroendocrinology, that the study of the control of the more metabolic functions in insects has lagged behind by several decades. I t was really only in the past ten years that metabolic hormones in insects, which all turned out to be neurohor mones, were seriously studied, and the real success stories from the point of view of the endocrinologist, the isolation, identification, and synthesis of such neurohormones, have broken within the past five years.
We have learned very recently that insects also possess insulin- or glucagon-like hormones (Chap. 5). However, in retrospect, the existence of specific metabolic neurohormones should have been expected in inver tebrates with no less certainty than is now known for vertebrates. This only shows that the dogma, still ripe when I did my first endocrinological studies with insects, that hormones were something special for ver-
Foreword and Overview XI
tebrates and developed very late in animal evolution, took a long time to die.
The general concept of neurosecretion and neurohormones hardly goes back 40 years and was crystallized largely in the work of the Scharrers.2
But this was foreshadowed in the early 1920's by Kopec's discovery of the brain function in the development of Lepidoptera, which took almost 30 years to be recognized as the driving force in insect development. Al though the role ofthe "brain" hormone, as it was first called, was well es tablished in the early 1950's and investigations and speculations on the nature of what is now most often called the prothoracicotropic hormone (Chap. 11) followed each other in an unending stream, we are now, 30 years later, still very largely in the dark about the identity ofthis hormone, as the last chapter in this book surprisingly reveals.
The ways in which scientific concepts develop are often strange and devious, and nothing illustrates this better than the topic of the hormonal control oftanning in insects, a subject I have been connected with, on and off, for over 45 years and which came to play also a dominant role in the development of our concepts in insect neuroendocrinology. The hormone now known as ecdysone, was originally discovered as the factor that brings about tanning of the fly puparium. Although the wider implication of ecdysone in molting and metamorphosis was soon recognized. it took over 25 years to recognize that in pupariation, ecdysone controlled not only tanning but also other morphogenetic events that bring about pupariation, though only indirectly as it turned out later. The all impor tant role of ecdysone as the tanning hormone was generally assumed for 30 years, when another hormone, bursicon, a product of neurosecretion (Chap. 6), was recognized as the tanning hormone for the adult fly. It then turned out that the role of ecdysone in tanning of the pupariation was an exception, a freak, as it were, among insects, and that possibly all conven tional tanning after a molt is generally controlled by bursicon. Surpris ingly now, even the concept of ecdysone as the tanning hormone in pupariation no longer seems to be true, as follows from the discovery of the pupariation factors (Chap. 7), neurohormones which are set in motion by ecdysone, one of which (PTF) seems specifically to have the function of controlling tanning.
Bursicon, which originally was found just to effect tanning is now seen also to control many other events during the consolidation of the cuticle after a molt, plasticization during general, and specifically wing expan sion, deposition of the endocuticle, cell death between the lamina of the wings, and possible formation of the apodemes. Fortunately, this does
2 Scharrer, E., Scharrer B. (1963). Neuroendocrinology. New York. Columbia University Press. 289 pp.
XII Foreword and Overview
not invalidate the propriety of the term, which was originally derived from the Greek bursicos-pertaining to tanning-because this is derived from the word bursa-skin. So bursicon now stands appropriately as a term for a hormone that gives the insect cuticle its peculiar properties after a molt.
The history of insect endocrinology, and particularly neuroen docrinology, is replete with surprising discoveries that uncovered the ex istence of unique processes or adaptations. These discoveries could only have been made originally by observers familiar with good, "old fashioned" natural history. It is as if "nature" had contrived to reveal its secrets to the observer in certain rare and striking phenomena. Let us consider a few notable examples.
The adipokinetic hormone (Chap. 2). Locusts use fat as energy for flight, in contrast to many other insects which use carbohydrates. The fat is stored in the fatbody and released into the hemolymph within a few minutes of beginning of flight.
Bursicon and plasticization hormone in flies (Chaps. 6 and 8). The adult fly emerges from the puparium in the soil and has to dig its way out before it expands body and wings and tans the body. These processes, to be effective, must be delayed (inhibited) until the fly is free from the soil. Then they are initiated by bursicon, which plasticizes the cuticle to make it inflatable, then tans the body, and subsequently controls a number of other processes. Similar processes are operating in other insects, but it was the particular ease with which they can be demonstrated and tested in flies which at first led to these discoveries.
Plasticization and diuretic hormones in Rhodnius (Chap. 8). At the very beginning of insect endocrinology stands the discovery that Rhod nius, a then obscure South American large blood sucking bug, takes only one blood meal in each instar. This blood meal can be 12 times the vol ume of the body, and this is only made possible by the secretion of the plasticization hormone which makes the cuticle expandable. Sub sequently, the diuretic hormone is released which controls the rapid excretion of the excess water in the blood. Similar events probably occur in other blood-sucking insects.
The Pupariation factors (Chap. 7). Puparium formation in flies (pupariation) is a unique morphogenetic event among insects and has proved of enormous heuristic value in insect endocrinology. In this pro cess, a soft, colorless larva contracts into a rigid dark puparium under the influence of what is now recognized as a series of hormonal events. One of the beauties of these reactions is that they take place within one hour. It started with the discovery of the hormone now known as ecdysone. Thirty-five years later the pupariation factors (Chap. 7) were discovered, neurohormones set in motion by ecdysone that control a variety of
Foreword and Overview XIII
manifestations during pupariation, anterior retraction (ART), immobiliza tion (PIF), possibly a stimulation factor (PSF), and ultimately tanning (PFT). It is still not known whether neurohormones like the pupariation factors are unique in this process, or are elicited by ecdysone in also other contexts.
The eclosion hormone (Chap. 9). Recognition that eclosion from a pupa is controlled by a specific hormone is of very recent date, and still confined to a few species of moths. This hormone triggers typical eclosion behavior even in an isolated abdomen!
Diapause hormone in Bombyx mori (Chap. 10). Diapause (arrest of development) occurs in many insects in a great variety of manifestations, and is often caused by a lack of ecdysone. But the recognition of a specif ic diapause hormone in the common silkworm is so far unique. This was the outcome of an enormous and prolonged effort to breed different races of silkworms in Japan.
Making good use of the specific reactions that led to the discovery of the various insect neurohormones, the following, mostly rapid and specif ic tests were developed:
Proctolin: Motility of the isolated cockroach hindgut (proctodeum). Adipokinetic hormone: Mobilization of lipids from the locust fatbody,
in vivo and vitro. Diuretic hormone: Elimination of fluid from isolated Malpighian
tubules of Rhodnius. Bursicon: Neck ligation in a fly immediately after emergence, tested for
tanning. Other tests proved less specific and convenient. Pupariation factors: Acceleration of pupariation and tanning in Sar
cophaga larvae selected several hours before pupariation (early red spiracle larvae).
Cuticle pLasticizing factors: Stretchability of cuticle in neckligated flies immediately after emergence (as in bursicon test), or stretchability of Rhodnius cuticle immediately after a blood meal.
Eclosion hormone: Precocious eclosion of the pharate adults of Antheraea pernyi; or induction of eclosion behavior in ligated abdomens of HyaLophora cecropia several hours before natural eclosion.
Diapause hormone: Injection of brain-suboesophageal ganglion ex tracts into pharate adults of non-diapausing strains of Bombyx mori. An important feature of this test is the fact that diapausing eggs are colored.
Prothoracicotropic hormone: The brains of the Satumiid Samia cynthia ricini were removed early in the pupa. The test consisted of in ducing adult development, and proved superior to, and more reliable than, previous attempts with Bombyx mori and H. cecropia pupal assays, or a larval assay with Manduca sexta.
XIV Foreword and Overview
The existence of unique processes in insects which led to the discovery of the many hormonal reactions, and the resulting opportunity to turn these situations into sensitive and rapid tests tended to offset the difficulty of using insects in hormonal research, inherent in their small size. The one situation where this becomes a serious obstacle is when it comes to isolation and identification. The number of individual insects which have been collected or worked up in particular tests boggles the mind. To give here a few examples:
Proctolin: 180 JLg were isolated from 125 kg of cockroaches, appro 125,000 individuals.
Adipokinetic hormone: Isolated from "only" 3000 corpora cardiaca in dividually dissected out from locust heads.
Diuretic hormone: The collection of material presented a major problem. One-hundred ganglionic masses can be collected from Rhodnius in one hour, but several thousand are used in an experiment.
Eclosion hormone: 300 g of eyeless heads from 6000 Manduca sexta adults were worked up.
Diapause hormone: In one experiment, the heads of one million male Bombyx mori moths were collected, yielding 2 kg of powder. In an earlier attempt the suboesophageal ganglion-brain complexes were dissected from 15,000 pupae.
Prothoracicotropic hormone: Working up 100,000 pupal brains did not prove enough material. Later whole heads were used with greater suc cess, working them up in batches of 48,000 isolated heads.
How far have we actually come in learning about the chemical nature of the various neurohormones? Only in two cases, appropriately described in the first two chapters of this book, has isolation proceeded to full iden tification, and that was achieved only within the past several years, doubtlessly made possible by the enoromous progress in chemical tech nology.
Proctolin is a pentapeptide of the formula H-Arg-Tyr-Leu-Pro-Thr OH, and the adipokinetic hormone (AKH) is a blocked decapeptide with the formula PC A-Leu-Asn-Phe-Thr-Pro-Asn-Trp-G ly-Thr-N H 2•
So far, all insect neurohormones have been shown to be of polypeptide or protein nature, and to be inactivated by proteases. On less advanced levels of isolation the approximate molecular weights (MW) have been stated with a greater or lesser degree of accuracy. A molecular weight of about 40,000 for burs icon has been confirmed several times. Among the pupariation factors, ARF, about 180,000, is possibly identical with PIF; and PTF is about 320,000. The cuticle plasticizing factor is probably identical with bursicon, with a molecular weight between 30,000 and 60,000.
The eclosion hormone, about 9,000, is tentatively considered a polypeptide of about 70 amino acids in length. Two active fractions of
Foreword and Overview XV
MW of 3,300 and 2,000 respectively, the former containing 14 kinds of amino acids and 2 kinds of amino sugars, and no sulfur-containing amino aci5is are reported for the diapause hormone. The latest figures for prothoracicotropic (brain) hormone show a MW of 4,400 daltons, which still have to be reconciled with earlier claims of 5,000, 9,000, 12,000, 20,000 and 31,000 daltons.
With this multiplicity of claims concerning the "brain" hormone (PITH), one wonders whether the end ofthe road has now been reached, or whether this elusive hormone may not in the end turn out to be of mul tiple nature, or of different nature in different insects. In fact, the search for the "brain" hormone has turned out to be a veritable exercise in frus tration, due undoubtedly, to the absence of precise, rapid and reliable test ing methods. A number of adverse factors have combined here to make these undertakings a misery, such as interference by photoperiodic phe nomena, responses to unspecific materials, such as metallic ions or cholesterol, the relatively long time of response, difficulty in raising the test animals, and the complexity of surgical procedures.
I cannot close this preface without at least mentioning three gaps, ex plainable by the emphasis laid in this book on the hard facts of testing, isolation and identification, but still of great relevancy to the subject.
The merits of Manfred Gersch and his school in Jena, Germany, in focusing on the role of neuroendocrinological events in development and metabolism over a period of 25 years have not been fully appreciated.
The role of allatotropic hormones, which mobilize the juvenile hor mone, has been postulated or claimed ever since the parallel activity of the prothoracicotropic hormone was recognized. Evidence for such hor mones seems overwhelming, both in the general control of the molt and metamorphosis and in the more specific control of oogenesis and yolk deposition, but no attempts of isolation or characterization seem to have ever been forthcoming.
And last but not least, the book fails to convey an appreciation of the morphological-neuroanatomical basis of all our knowledge in the field, largely based on the conceptual work of the Scharrers,2 and as far as in sects are concerned, the invaluable contributions of Bertha Scharrer over more than 40 years.
Department of Entomology University of Illinois Urbana, Illinois 61801 February 1980
GOTTFRIED S. FRAENKEL
Contents
Chapter 1 Proctolin: Bioassay, Isolation, and Structure A.N. STARRATT and R.W. STEELE. With 4 Figures
I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. 1 II. Bioassay .................................................. 3 III. Chemistry ............................................... 13
References .............................................. 28
Chapter 2 Adipokinetic Hormone Judith V. STONE and W. MORDUE. With 18 Figures
I. Introduction ............................................. 31 II. Biological (Bioassay) ...................................... 33 II I. Chemical ................................................ 45
References .............................................. 76
Chapter 3 Bioassay of Diuretic Hormone in Rhodnius S.H.P. MADDRELL. With 5 Figures
I. Introduction ............................................. 81 II. Isolation of Malpighian Tubules from Rhodnius ............. 82
References .............................................. 90
XVIII Contents
Chapter 4 Diuretic Hormone-Extraction and Chemical Properties RJ. ASTON and L. HUGHES. With 6 Figures
I. Introduction .............................................. 91 II. Assay of Hormone Activity ............... , ............ " ... 93 III. Isolation of Diuretic Hormone Storage Tissue ................ 94 IV. Methods of Fractionation .................................. 95 V. High K + Release of Diuretic Hormone In Vitro ............. 107 VI. Properties ............................................... 108 VII. Cross-Reactivity of Insect Diuretic Hormones .............. 111 VIII. Concluding Remarks ......................... , ............ 111
Acknowledgements ....................................... 112 References ............................................... 112
Chapter 5 Insulin-like and Glucagon-like Hormones in Insects K.J. KRAMER. With 4 Figures
I. Introduction ............................................. 116 II. Preparation of Tissue Extract. ............................. 117 II I. Purification of Ex tract and Heterogeneity ................... 119 IV. Biological Assay .......................................... 124 V. Radioimmunoassay ....................................... 127 VI. Immunocytochemistry .................................... 131 V I I. Concluding Remarks ..................................... 13 I
Acknowledgement ........................................ 132 References .............................................. 132 Note Added in Proof ..................................... 136
Chapter 6 Bursicon I.M. SELIGMAN
I. Introduction ............................................. 137 II. Purification of Bursicon ................................... 144 III. Assays for Bursicon Activity ....................... , ....... 145
References ............................................... 150
Chapter 7 Neurohormonal Factors Involved in the Control of Pupariation J. ZbAREK. With II Figures
I. What are the Pupariation Factors? ......................... 154 II. Choice of Material ...................................... .156
Contents XIX
III. Breeding Technique ....................................... 156 IV. Staging of the Larvae for Experiments on Pupariation ........ 157 V. Methods of Observing and Recording Pupariation ............ 158 VI. Bioassays for the Activity of the Pupariation Factors ......... 164 VII. Materials Possessing Activity of the Pupariation Factors ..... 169 VIII. Chemical Identification of the Pupariation Factors ........... 170 IX. Remarks to the Mode of Action of the Pupariation Factors ... 175
References ............................................... 177
Chapter 8 Cuticle Plasticizing Factors S.E. REYNOLDS. With 5 Figures
I. Introduction .............................................. 179 II. Bioassay ................................................. 183 III. Chemistry ................................................ 188
References ............................................... 193
Chapter 9 Eclosion Hormones S.E. REYNOLDS and J.W. TRUMAN. With 10 Figures
I. Introduction .............................................. 196 I I. Bioassay ................................................. 199 III. Chemistry: Isolation and Purification ....................... 205 IV. Properties ................................................ 210 V. Biological Activity of the Purified Hormone ................. 2\3
Acknowledgements ....................................... 214 References ............................................... 214
Chapter 10 Diapause Hormones M. (SOBE and T. GOTO. With 19 Figures
I. Introduction .............................................. 216 II. Materials ................................................. 218 III. Bioassay ................................................. 221 IV. Extraction ............................................... 223 V. Chromatographic Separation ............................... 225 VI. Selective Extraction ....................................... 227 VII. Chromatography on Merckogel OR 6000 ................... 231 VIII. Isolation of DH-A and DH-B .............................. 232 IX. Molecular Weight ......................................... 233 X. Stability of DH in Relation to Degree of Purity .............. 234 XI. Activity of the Two Species ................................ 235
:xx Contents
XII. Stability and Characters .................................. 237 XIII. Infrared Spectra ......................................... 239 XIV. Constituents ............................................. 240
Acknowledgments ....................................... 241 References .............................................. 24 I
Chapter 11 Prothoracicotropic Hormone H. ISHIZAKI and A. SUZUKI. With 8 Figures
I. Introduction ............................................. 244 I I. What is Known?-Biological. .............................. 245 III. Bioassay ................................................ 249 IV. Chemistry ............................................... 256
References .............................................. 271
Index ........................................................ 277
S. E. Reynolds
List of Contributors
ARC Unit of Invertebrate Chemistry and Physi ology, The University of Sussex, Brighton BN I 9QJ, England Laboratory of Organic Chemistry, Faculty of Agriculture, Nagoya University, Nagoya 464, Japan ARC Unit of Invertebrate Chemistry and Physi ology, The University of Sussex, Brighton BN 1 9QJ, England Biological Institute, Faculty of Science, Nagoya University, Chikusa-ku, Nagoya 464, Japan Laboratory of Organic Chemistry, Faculty of Agriculture, Nagoya University, Nagoya 464, Japan USDA, Grain Marketing Research Laboratory, 1515 College Avenue, Manhattan, Kansas 66502, USA Department of Zoology, University of Cam bridge, Cambridge CB2 3EJ, England Department of Zoology, University of Aber deen, Aberdeen AB9 2TN, Scotland Animal Physiology and Ecology Group, School of Biological Sciences, University of Bath, Claverton Down, Bath BA2 7 A Y, England
XXII List of Contributors
J. Zdarek
Institute of Developmental Biology, Texas A & M University, College Station, Texas 77843, USA Research Institute, Agriculture Canada, Univer sity Sub Post Office, London, Ontario N6A 5B7, Canada Research Institute, Agriculture Canada, U niver sity Sub Post Office, London, Ontario N6A 5B7, Canada Department of Zoology, Imperial College of Science and Technology, Prince Consort Road, London SW7 2AZ, England Department of Agricultural Chemistry, Univer sity of Tokyo, Bunkyo-ku, Tokyo 113, Japan Department of Zoology, University of Washing ton, Seattle, Washington 98195, USA Institute of Entomology Csav, Department of Insect Physiology, Papirenska 25, Praha 6, Czechoslovakia
ACTH AKH ARF AZT CA c-AMP CC CNS DDSA DEAE DFP DH DH-A DH-B DHE DMP DOPA OTT H.p.l.c. 5-HT JH KIU aMDH
MH
XXIV
MTGN NADA NBS NEM 3-0HK PDF PIF PNC PSF PTF PTTH RIA RPCH SDS SG TCA VeIVo
List of Abbreviations
Chapter 1
A. N. Starratt and R. W. Steele
I. Introduction
More than a decade ago Brown (1967) reported the extraction of a myo tropic substance from the viscera of the cockroach, Periplaneta ameri cana (L.), and proposed that it might function as an excitatory neurotrans mitter in the visceral muscles of insects. This "gut-factor," later called proctolin, caused slow-type graded contractions of the longitudinal muscles ofthe hindgut (proctodeum) similar to those evoked by repetitive nerve stimulation. Pharmacologically, it differed from any of the known or suspected neurotransmitters tested, including 5-hydroxytryptamine, acetylcholine, adrenaline, noradrenaline, y-aminobutyric acid, and glu tamic acid. It was also different from two peptides that have activity on the hindgut, which Brown (1965) isolated from extracts of P. americana corpus cardiaca.
After considerable effort, proctolin was isolated (Brown and Starratt 1975) and was identified as the pentapeptide H-Arg-Tyr-Leu-Pro-Thr OH (Starratt and Brown 1975). To confirm this structure, the peptide that has this sequence was synthesized (Starratt and Brown 1977). Chromatographically, electrophoretically, and pharmacologically, the synthetic peptide was identical to natural proctolin.
A survey of representatives of six insect orders has indicated that proctolin is widely distributed (Brown 1977). Each of the eight species examined yielded a substance with myotropic activity on the cockroach hindgut when extracted and partially purified by using a modification of the method employed for the isolation of proctolin. Pharmacological,
2 A.N. Starratt and R.W. Steele
chromatographic, and electrophoretic properties of the substance from each of the species were identical to those of proctolin. Although identity was not unambiguously established and only a relatively few species were examined, these results led Brown (1977) to propose that proctolin may be a universal constituent of the Insecta. Similarities between proctolin and a hindgut-stimulating peptide that Holman and Cook (1972) obtained from hindguts, terminal ganglia, proctodeal nerves, and heads of another cockroach, Leucophaea maderae, resulted in the suggestion that these peptides were probablY identical (Brown and Starratt 1975; Starratt and Brown 1975). However, although Holman and Cook (1972) also found their hindgut-stimulating peptide in P. americana and the grasshopper, Schistocerca nitens, they were unable to detect it in foreguts from L. maderae, in the head of the housefly, Musca domestica, or in fifth-instar larvae of the tobacco homworm, Manduca sexta. Moreover, Holman and Cook ( 1972) suggested that their peptide acted as a neurohormone in volved in modulating muscle excitability and supported their hypothesis in subsequent papers (Cook and Holman 1975; Cook et al. 1975). By contrast, in a later paper Brown (1975) presented additional data consis tent with his earlier proposal that proctolin acted as an excitatory trans mitter (Brown 1967). These differences point out the need for further studies to determine the distribution of proctolin in the Insecta and to es tablish its physiological role as a neurotransmitter or neurohormone.
In addition to causing contractions of the slow striated muscles of the gut of P. americana, proctolin has been found to be active on other insect muscles and nerves: It induces myogenic contractions in a leg muscle of two species oflocust at concentrations of 10-10 to 10-9 M (Piek and Man tei 1977; May et a1. 1979), it increases the rate and amplitude of contrac tion of semi-isolated heart preparations from P. americana at a threshold concentration of about 10-9 M (Miller 1979), and it increases nervous ac tivity when assayed on the ventral nerve cord supply to the hypemeural muscle of P. americana (Miller 1979).
A number of studies have indicated several peptides in the insect ner vous system that act on the gut and affect the heartbeat rate of insects (Frontali and Gainer 1977). The isolation and characterization of these substances present a challenge that must be met before it can be known if any structural similarity exists between these peptides and proctolin, the first insect neuropeptide to be identified.
A description of the methods utilized and found to be satisfactory for the detection, extraction, isolation, and characterization of proctolin is presented in this chapter. In general, the sequence of steps is the same as would be followed for the isolation and identification of any physiologi cally active peptide. Probably, the availability of a facile and reliable bioassay procedure was the most important factor contributing to the success of this work with proctolin. It is hoped that this account will be
Proctolin: Bioassay, Isolation, and Structure 3
useful as a guide to anyone undertaking investigations of proctolin or other peptides, especially those exhibiting physiological activity on insect viscera.
II. Bioassay
The neuropharmacological procedures described in this section were developed to show the action of proctolin on the longitudinal muscles of the whole proctodeum of adult male P. americana, to demonstrate the physiological effects of stimulating the proctodeal nerves, and finally, to compare the interactions of nervous stimulation with proctolin and other agonists and antagonists. The technique is sensitive, convenient, and relatively rapid. The whole proctodeum with its intact nerve supply is isolated and suspended for isotonic recording in a suitable organ bath of known volume. Proctolin and other drugs are added to the bath and their effects on the proctodeum are examined with reference to the responses evoked by nerve stimulation. In addition, simple bioassays of proctolin activity may be carried out in a delightfully straightforward manner since the proctodeum without its nerve supply can be isolated and suspended for assay in a few minutes. The methodologies involved are those com monly used in physiology and pharmacology (for example, Staff 1970).
A. Isolation of the Proctodeum
As in many insects, the cockroach proctodeum or hindgut is divided by a constriction into two regions, the anterior intestine and the posterior in testine or rectum. A full description of the musculature and innervation ofthese regions is given by Brown and Nagai (1969). To summarize, both regions possess circular and longitudinal muscles, but the organization of fibers in the two regions is substantially different. In the anterior intes tine, longitudinal muscles are organized into many short, flat bundles that intimately associate with the mainly underlying, circular muscle fibers. On the rectum, the longitudinal muscle fibers are limited to six discrete bundles, symmetrically placed around the anterior two-thirds of the rec tum. Each bundle consists of independent inferior and superior straps, which overlie a thin layer of circular muscle that is considerably thickened at the intestinal constriction and around the anus. Six fan shaped bundles of rectum dilator muslces are inserted on the posterior third of the rectum. These dilator muscles originate from the anterior edges of the 10th abdominal sclerites, with the dorsal and lateral pairs of dilator muscles from the 10th tergite, and the ventral dilator pair from the 10th stemite.
The proctodeum is innervated by the proctodeal nerves, bilateral dorso-medially directed branches of the cereal nerve XI (Roeder et al. 1960). Shortly after branching from the cereal nerve XI, the proctodeal nerve divides into an anterior and posterior branch, although occasionally these branches emerge separately. The posterior proctodeal nerve inner vates the dorsal and lateral rectum dilator muscles, and the circular muscle of the posterior region of the rectum. The anterior branch of the proctodeal nerve supplies the rectum longitudinal muscles, the ventral dilator muscles, the circular muscles of the anterior region of the rectum, and all the muscles of the anterior intestine. The latter muscles are inner vated by four major nerve trunks that originate by division of each an terior proctodeal nerve in the region ofthe intestinal constriction. Beside these components of central innervation is a suggestion that some form of peripheral innervation is localized to muscles in the region of the rectal valve, although proof of such peripheral innervation is lacking (Brown 1975). Some of these proctodeal components and their in situ rela tionships can be seen in Fig. 1-1.
Vth abdominal ganglion
9th abdominal tergite --___ 'Oth abdominal tergite
(supraanal plate)
The muscle fibers of the proctodeum undergo coordinated contractions that give rise to peristalsis. Although it is difficult to follow the behavior of the circular muscles, that of the longitudinal muscles can be recorded easily and provide the subject for neuropharmacological assay. The isolation procedures described below differ in certain respects from the methods used by Holman and Cook (1970). These differences may only be trivial, but any interested investigator is urged to attempt both procedures to determine the best for his own use.
Before dissection, immobilize the male cockroach by briefly chilling it on ice. Males generally are easier to dissect than are females because their reproductive system is more discrete and, thus, far easier to dissect
(b)
Figure 1-1. Isolation of the proctodeum from the abdomen of a male P. americana . (a) Drawing of the dissected abdomen prior to cutting the visceral tracheal trunks on the left side as described in the text. Major anatomical features are noted. (b) In situ view after dissection of these tracheae.
without damage to the fine proctodeal nerves. Remove the legs and wings, and pin the specimen through the metathoracic coxites, dorsal side up, in a dissecting tray. Flood the tray with fresh insect saline containing 9.0 g NaCl, 0.2 g KCI, 0.2 g CaCI2 , 3.96 g dextrose, and 10 ml 0.1 M sodium phosphate buffer, pH 7.0, per liter (after Pringle 1938). The pH of this Ringer's solution is 6.9 and is unchanged by oxygenation (Brown 1965). With fine dissecting scissors make a superficial midline incision through the last abdominal sclerites (7th and concealed 8th tergite) and continue through to the thorax. Gently open the dorsal surface and pin it aside to expose the viscera. Ideally, the specimen should be stretched slightly in its long diQ1ension and pinned so that the nerve cord connec tives lie flat on the floor of the abdomen.
Carefully cut the trachea and Malpighian tubules that invest the surface of the anterior intestine. Once freed of these restraints, the proctodeum is severed just anterior to the point of insertion of the Malpighian tubules and is placed to one side. The remaining alimentary organs and much of the fat body can now be cleared from the abdominal cavity to expose the ventral nerve cord. Next, the accessory glands are removed by pulling these organs dorso-anteriorly and proximally severing their gonophore connections. With this accomplished the large cercal nerves will be visi ble as flat bundles emerging from the Vlth abdominal ganglion, and the proctodeal nerves can be traced from the cereal nerve XI (see Fig. 1-1 a).
Take up the supraanal plate or 10th tergite (Snodgrass 1937) and dis sect along the midline towards the anterior. While still lifting the 10th tergite, continue the incision through the concealed 9th tergite and its in tersegmental membrane, but avoid damage to the underlying posterior region of the rectum. Each half of the freed 9th tergite can now be pulled to one side with forceps, thereby displaying the fan-shaped bundle of dor sal rectum dilator muscles (Fig. 1-1). Working on one side, carefully cut these muscles close to their points of origin on the anterior edge of the 10th abdominal tergite. This procedure effectively reveals the paraprocts and central lobe-shaped epiprocts that lie immediately beneath the margin of the 10th tergum. Cut away the membraneous attachment between these sclerites and the 10th tergite, continuing the cut around the dorsal edge of the papaproct and into the membranous socket of the cercus. This frees the cercus from the 10th tergum, and it also severs the fan shaped bundle of lateral rectum dilator muscles that originate on the ex treme lateral margin of the 10th tergite. The cercus and tergites on this side can now be pinned aside, as in Fig. 1-1. If dissected correctly, the cercal nerves X and XI remain attached to the lateral margin of the ex posed paraproct. This feature, although not strictly necessary for a suc cessful dissection, greatly simplifies the later ligaturing of these nerves and also serves to keep the proctodeal nerves clear during dissection of the ventral rectum dilator muscles. Continue the dissection on the ex-
posed side by taking up the visceral tracheal trunks that supply the rectum (Fig. 1-1 a). Stretch each trunk dorso-Iaterally until it is clear of the cer cal and proctodeal nerves below and then cut. Note that both the anterior and posterior branches. of the proctodeal nerve carry a fine tracheol along their length, and great care must be taken not to damage these nerves dur ing this step. Repeat these procedures on the other side. Should the ex perimental design require only pharmacological assay, ignore the latter precautions and sever all connections between the proctodeum and its nerve supply.
Next, lift the paraprocts and cut their ventral membranous connection to the genital pouch that contains the asymmetrical, hooked phallomeres. The proctodeum now lies free in the abdominal cavity except for the proc todeal nerves branching to the cercal nerves XI (which emerge from the Vlth abdominal ganglion, Fig. 1-1), some minor attachments to the 10th abdominal sternite by degenerated intersegmental muscles of the 10th and 11th segments (Brown and Nagai 1969), and two strong fan-shaped bundles of ventral rectum dilator muscles also attached to the 10th ab dominal sternite. To continue the isolation, take up the papaprocts in for ceps and gently lift anteriorly. The cercal nerves, if still attached to the lat eral margins of the paraprocts, will lift clear and allow the remaining at tachments to the 10th sternite to be ventrally severed without damage to the proctodeal nerves. Continue to work forward and free the last two ab dominal ganglia from their tracheal and peripheral neural attachments, then sever the nerve cord near the Vth abdominal ganglion. The isolated proctodeum with its intact nerve supply can now be removed from the ab domen and placed in a petri dish that contains oxygenated saline. Preparations destined solely for pharmacologic assay require only the severence of the ventral rectum dilator muscles for complete isolation.
At this point it is well to observe the preparation. If the nerve supply remains intact, the proctodeum should be undergoing spontaneous con tractions. Moreover, it should readily contract upon mechanical stimula tion since the dominant contractile elements, the six superior rectallongi tudinal muscle bundles, appear myogenic with central nervous control (Nagai and Brown ,1969). If these conditions obtain, the proctodeum is prepared for myographic recording as follows. First, the left and right cercal nerves are tied together to ensure later introduction of both proc todeal nerves into the stimulating suction electrode. To accomplish this task, a 3-4 cm length of silk thread is separated into its individual strands with forceps, and one strand then is further subdivided into quarters or groups of 4-8 fibers. Prepare a loop in one group of fibers, pass it over the remnant of nerve cord, and tie a ligature around the cercal nerves just after their emergence from the Vlth abdominal ganglion. With fine dis secting scissors, carefully trim any excess silk and subsequently remove the Vlth abdominal ganglion. Most spontaneous neurogenic activity
should cease after this step. Next tie a 5-10 cm silk thread to the rectal end of the proctodeum, ligaturing the thread around the epiprocts or around a lateral corner of a paraproct. Finally, prepare a third thread 15-25 cm long and ligature around the anterior intestine immediately pos terior to the point of insertion of the Malpighian tubules. Simple bioas says require only the latter ligatures to ready the preparation for suspen sion in the organ bath.
B. Bioassay Apparatus
The 4-ml organ bath illustrated in Fig. 1-2 was fabricated in this labora tory to provide a simple and convenient experimental setup for bioassay. I t consists of a glass tube open at both ends with a side arm, a strategically placed suction electrode fixed in the side wall, and a rubber stopper in the bottom. A 22-guage steel pin penetrates the stopper and this pin is fashioned into a hook to anchor the tie from the rectal end of the proc todeum. Adjacent to this pin are two steel tubes that penetrate the stop per, one a 2-mm internal diameter needle for perfusion of the bath with the same Ringer's solution as described in Sect. II.A, and the other a 30-gauge hypodermic needle for delivery of oxygen. The Luer-Iok fitting on the latter permits easy disconnection from the oxygen reservoir, a fea ture that facilitates handling operations during ligature attachment and mounting.
Take the free end of the thread attached to the epiprocts or paraproct and tie to the pin in the organ bath stopper. Knot the thread so that about 5 mm separates the epiprocts from the pin. In neuropharmacological assays, this distance is important for easy interposition of the ligatured cercal nerves into the fixed suction electrode; indeed, wide divergence from the correct length can result in the preparation being suspended by the delicate proctodeal nerves, a situation that must be avoided. Lay the organ bath in the petri dish and with the thread attached to the anterior in testine draw the proctodeum into the bath chamber. During this step it is useful to perfuse the preparation with saline delivered through the stop per. Seat the stopper firmly and then fill the organ bath with saline and mount it directly below the lever arm of an isotonic transducer arranged in a system so that both the organ bath and transducer can be moved up and down independentlY. A Narishige MD-2 micromanipulator makes a par ticularly stable and flexible lower stage for mounting the organ bath.
Complete the suspension of the proctodeum by attaching the free thread from the anterior intestine to the lever of the transducer and apply a low tension of:o;;;; 50 mg to stretch the preparation. The transducer lever should be carefully balanced before the proctodeum is connected to it. Rotate and/or elevate the anchor pin in the stopper so that the ligatured
Suction electrode
isolation unit
Miniature clip --~
i---Glass organ bath
tr-+-"'--- To buffer reservoir
Figure 1-2. Diagram of the organ bath assembly used for proctodeal bioassay. The glass bath chamber is 60 mm x II mm internal diameter, with a glass side arm 50 mm x 7 mm internal diameter. The fixed suction electrode is 1.25-mm glass tubing drawn to a tip of 0.3 mm lumen and cemented with araldite in the side wall 12 mm from the chamber base. Stimulating and indifferent electrodes are platinum wire , 0.1 mm diameter, soldered to copper wire terminals of about I-mm diameter. Suction is provided by a 10-ml syringe.
cercal nerves align with the tip of the suction electrode. To obtain good electrical contact for stimulation, it is necessary that as little shorting as possible occurs between the suction electrode and the indifferent elec trode (Lang 1972). This is achieved by having the smallest tip opening that will accomodate the cercal nerves without damaging them. A tip with a lumen size of 0.3 mm has proved satisfactory. Provide suction using a 5-10 ml syringe attached to the suction electrode and gently draw up the cut ends of the ligatured cercal nerves. Observe the progress under a microscope and make sure that enough saline is in the electrode tip to
bridge the distance between the cercal nerves and the platinum electrode. Both the cercal nerves and proctodeal nerves will be lifted away from the freely suspended proctodeum, eliminating possible damage during con tractions. If all appears in order, the tension on the preparation can now be increased to the recording value of 200--250 mg, and oxygenation of the bath can commence. Adjust the oxygen flow to achieve a gentle stream of bubbles. A simple setup such as a bypass line with a pinch-cock will serve adequately, but superior control is obtained by using a supplementary valve such as a Nupro B-2SGD fine metering valve in the oxygen supply line.
The proctodeum requires a minimum of 60-90 min under 250-mg ten sion before the longitudinal muscles relax to constant length. A con siderable increase in sensitivity to proctolin and/or neural stimulation ac companies this stretching and assays should not be performed until relax ation is complete; thereafter, the preparation remains viable for 8-12 h which permits the assay of many samples.
In our experimental system, the platinum leads from the suction elec trode are connected in the conventional manner to a Grass SIU-5 stimulus isolation unit coupled to a Grass S-88 stimulator. Proctodeal contractions are recorded isotonically with a Harvard Model 386 trans ducer connected to a Harvard Model 350 recording module and a Havard Model 485 chart recorder. Event/time records are provided by a Harvard Model 284 module. Other stimulators, transducers, and/or recorders may be used; the choice of equipment depends solely on the laboratory facilities available.
c. Bioassay Procedures
Proctolin and other agonists are added rapidly to the organ bath in 5-200 JLI quantities delivered via Hamilton syringes. The drugs, which are made up in Ringer's solution, are squirted from the submerged needle toward the stopper where oxygen bubbles aid the mixing process. After a chosen contact interval (usually 20 s), fast perfusion of the bath at 50-100 ml/min is initiated by releasing a hemostat clamping the 0.64-cm rubber inflow tubing from a 5-liter Mariotte flask reservoir of fresh Ringer's solution. The contractions evoked by proctolin contact should be rapidly followed by complete relaxation to the previously stabilized baseline level (see Fig. 1-3). After relaxation, perfusion can be returned to 5-10 ml/min to con serve buffer, and then stopped at 2-4 min in preparation for exposure to the next dosage. For quantitative data it is important to maintain equal contact time, consistent perfusion rates, and equal time between doses. These requirements can be achieved without difficulty using a hemostat
6
4
20
(a)
(b)
Figure 1-3. Some typical responses of proctodeal muscle preparations. Contrac tions were recorded isotonically under 250 mg tension as described in the text. (a) Graded responses to frequency of nerve stimulation at 2-20 Hz and 0.1 V, applied in 2-s trains at 30-s intervals. With this preparation, five repetitive stimu lations at 10Hz were required to overcome adaptation to 20 Hz. (b) Graded re sponses to nerve stimulation and proctolin applied for 20 s. Relaxation to the baseline tension was achieved by buffer perfusion at 96 ml/min.
and stopwatch, but for convenience and reliability the perfusion line from the reservoir has been split into two branches and two solenoid valves (Ascolectric, No. 8262C 103) separately programmed by a Chron Trol timer (Lindberg Enterprises, San Diego, CA) were incorporated to yield repetitive perfusion cycles automatically. Results are estimated in the usual fashion by comparing the responses of unknown proctolin doses against a log dose-response curve established with known proctolin standards that are applied in a Latin square.
In neuropharmacological assays, the forces of neurally evoked con tractions depend in large measure on the quality of electrical contact with the proctodeal nerves. Under good conditions, stimulation at 0.1-0.3 V gives satisfactory responses; poor contact requires a higher stimulating voltage. Nerve stimulation is supramaximal with a single stimulus of 0.5-ms duration. The contractions evoked by repetitive nerve stimulation are graded responses to frequency of nerve stimulation. Routinely, neurally evoked contractions of the proctodeum can reflect a difference of one impulse per second in the range 6-15 Hz applied in 2-3 s trains every 30 s (Fig. 1-3). Maximum responses occur at about 50 Hz. For detailed descriptions of the bioelectrics underlying the mechanical activity of rec tum longitudinal muscle fibers, Belton and Brown (1969), Brown and Nagai (1969), Nagai and Brown (1969), and Nagai (1970, 1972, 1973) may be consulted.
Three neuropharmacological criteria have emerged that together ap pear characteristic of proctolin action on the cockroach proctodeum. These criteria are as follows: Proctolin above a threshold concentration of about 10-9 M evokes sustained slow-type, graded contractions in the longitudinal muscles of the whole proctodeum; proctolin at subthreshold concentrations of 2-8 x 10- 10 M potentiates the graded responses to repetitive nerve stimulation; and proctolin-induced responses are sup pressed 70-90% by 15-s preincubation with tyramine at 2x 10- 6 M (Brown 1975). These criteria complement the patterns of sensitivity to enzymic hydrolysis that distinguish proctolin from two hindgut-stimulat ing peptides present in the corpora cardiaca of P. americana (Brown 1965). Thus, proctolin is resistant to hydrolysis by chymotrypsin (EC 3.4.4.5), trypsin (EC 3.4.4.4), carboxypeptidase A (EC 3.4.2.1), and car boxypeptidase B (EC 3.4.2.2), but it is readily inactivated by leucine aminopeptidase (EC 3.4.1.1). Incubation of a reaction mixture containing 200 ILl proctolin extract (100-200 ng proctolin), 40 ILl 0.5 M Tris buffer (pH 8.5), 10 ILl 0.125 M MgCI2, and 40 ILl leucine aminopeptidase (2 mg/ml in 0.02 M Tris buffer, pH 8.5) causes the loss of about 50% of proctolin activity after 8 min at 35°C (Brown and Starratt 1975; Brown 1977). These properties, in addition to the chromatographic and elec trophoretic characteristics described in the next section, should prove
useful in establishing the occurrence of proctolin in other tissues and organisms.
III. Chemistry
A. Extraction and Isolation of Proctolin
This section describes in detail a method for the isolation of proctolin from P. americana. The multistep procedure follows closely that reported by Brown and Starratt (1975), who obtained 180 ILg pure proctolin from 125 kg whole cockroaches. A shortened version of this procedure has been used by Brown (1977) to examine the occurrence of proctolin in a number of other insect species.
Early attempts to isolate sufficient proctolin for structure determination utilized gram quantities of excised viscera that contained relatively high levels of proctolin (Brown 1967). When it became evident sufficient ma terial could not be obtained by this approach, mainly because of the time required to remove the viscera, the methods were scaled up to accom modate kilogram quantities of whole cockroaches. Throughout the procedure, proctolin was determined by bioassay on the isolated hindgut of P. americana (see Sect. II). Quantitative results in terms of "rectum equivalents" were obtained by comparison of the intensities of contrac tions caused by unknowns with that caused by a standard extract of cockroach rectums. One rectum equivalent was defined as the amount of proctolin present in one rectum and is now known to be equal to about 0.86 ng proctolin. The bioassay provided a relatively facile means of monitoring the progress of the purification.
1. Extraction
All steps of the extraction were carried out in a cold room operated at 1°C to minimize the destruction of proctolin either chemically or, in the initial stage, enzymatically because of proteolytic enzymes present in the tis sues. The described procedure was that used during the "large-scale" isolation of proctolin, but it may be readily adapted for extraction of small quantities of cockroaches or other insects (Brown 1977).
In preparation for the extraction, cages containing the adult cockroaches were placed in the cold room. When immobile, the insects in groups of 1000 were transferred to a blender and homogenized thoroughly in 2 liter cold 7% perchloric acid. The thick suspension was filtered overnight under reduced pressure through cheesecloth and filter paper in a Buchner funnel. A 45% potassium hydroxide solution was
14 A.N. Starratt and RW. Steele
added with efficient stirring until pH 6 was reached. The pH was es timated by use of narrow-range indicator paper. After additional cooling, the precipitated potassium perchlorate was removed by filtration. The fil trates from 4000 insects were pooled and reduced in volume to about 15% of the original by rotary evaporation in vacuo at room temperature. After cooling of the concentrated filtrate to 1 °e, the clear solution was decanted from the precipitate that formed. This solution was held in a freezer at - 200e until extracts from additional batches of insects were accumulated. With smaller quantities of insects, the isolation proceeded without inter ruption (Brown 1977). Pooled extracts from 25 000 cockroaches were warmed to room temperature and diluted with an equal volume of ethanol. The precipitate formed was removed by centrifugation.
2. Isolation
The optimum purification scheme for any substance can be determined only by trial and error. Ideally, each step should be planned to provide the greatest increase in specific activity or purity and the best yield while requiring the minimum amount of time and effort. Although in the case of the large-scale multistep procedure used to isolate proctolin most of the fractionation methods were first tried by using small amounts of extract as well as reference amino acids and peptides, it cannot be claimed that the steps have been optimized or that the most efficient scheme has been found. Further studies with methods described here, as well as other separation methods such as high-performance liquid chromatography, will undoubtedly lead to improvements in the present scheme for the isolation of proctolin. However, in spite of the fact that all the work necessary to identify the best purification scheme was not performed, it is estimated that about 12% of the proctolin was recovered.
As well as providing guidance to those wishing to isolate proctolin from P. americana or other insects, the approach and methods described here may be useful to investigators trying to isolate other physiologically ac tive insect peptides. The steps used for the successful purification of proctolin are all standard and have been described in detail in reviews and books [for example, Morris and Morris (1976); Wolf (1969)] concerning separation methods used in biochemistry and organic chemistry. A few sources of information concerning particular methods are presented throughout this section. Since all investigators may not require a pure preparation of proctolin for their studies, the description of the isolation has been divided into eight steps. Each stage produced a preparation that evoked contractions of the cockroach hindgut. Table 1-1 summarizes the results. From the number of rectum equivalents present and the dry weights it was possible to determine specific activities. The quantity of proctolin shown for each stage was calculated after the final step. It is hoped that the table will be useful as a guide in determining the number of
Table 1-1. Scheme for purification of proctolin. Active
fraction Rectum Amt. Step Procedure (dry wt.) equivalents proctolin"
I Separation of extract from 125 000 cockroaches in 5 portions on Dowex 50W-x8(H+ form) and then on Dowex 50W-x8(NH4 +
form) 3400 mg 1.3 x 106 1.I2 mg
2 Alumina chromatography 660 mg 1.2 x 106 1.03 mg
3 Chromatography on Rexyn 101(NH4+ form) 240 mg 1.1 x 106 940 ILg
4 Craig countercurrent separation 35 mg 8.7 x 106 750 ILg
5 Paper chromatography 5.3 mg 7.0 x 105 600 ILg
6 Separation by high-voltage paper electrophoresis 4.7 x 105 400 ILg
7 Chromatography on Sephadex G-15b 2.4 x 105 206 ILg
8 Chromatography on Rexyn 101(NH4+ form) 180 ILg 2.1 x 105 180 ILg
'Calculated after final step and based on the weight and the activity of the pure proctolin (I rectum equivalent = 0.86 ng proctolin).
h75% of the active sample from Sect. III.A.2.f used.
steps necessary to provide a preparation containing proctolin sufficiently pure to meet the requirements of the work being undertaken.
a. Step I. As the initial purification procedure, the extract was passed through a column of Dowex 50W-x8(50-100 mesh, H+ form). Substitu tion of an equivalent ion-exchange resin of another manufacturer would be expected to produce similar results. A comprehensive discussion of ion-exchange chromatography covering theory, equipment, and tech niques is found in the book by Khym (1974), and an article by Schroeder (1972) provides a detailed description of a method for the separation of peptides on Dowex 50. The ion-exchange procedures described in this step served mainly to separate proctolin from substances chemically very different that constituted the bulk of the extract.
The ion-exchange resin was stirred with water and the fine particles were removed by decantation after a short settling period. Although not noted subsequently, all water used in this work was distilled. The resin was then stirred with 3 vol. (relative to the resin volume) 2 N hydrochloric acid. After standing for 30 min, the acid was poured off and the resin was washed three times with 3 vol. water each time. The resin
was then stirred with 3 vol. 2 N sodium hydroxide. After standing 30 min, the sodium hydroxide solution was decanted and the resin was washed three times with water. This procedure with acid and alkali was repeated twice. Finally, the resin was treated with 3 vol. 2 N hydro chloric acid and then was washed free of acid with water.
A 7 x 40 cm column of Dowex 50 (H+ form) was prepared by pouring a slurry of the resin into a chromatographic column equipped with a stop cock so that the flow could be interrupted when necessary. Water was passed through the column until the resin was fully settled. When ready to use, the aqueous ethanol solution (15 liter) containing the extract of 25 000 cockroaches was applied to the top of the resin bed and the column was eluted consecutively with 4 liters each water, I N pyridine, and a I: I mixture of 4 N ammonium hydroxide and ethanol. For smaller-scale runs such as those made during the investigation of the occurrence of proctolin in other species of insects, a 2.5 x 30 cm column and 300 ml of each of the eluates proved satisfactory (Brown 1977). After elution of the basic fraction, the column was regenerated by washing with 3 vol. each 2 N sodium hydroxide, water, and 2 N hydrochloric acid, followed by water until the washings were neutral.
By using a rotary evaporator, the solvent was removed at reduced pres sure (water aspirator) and room temperature from the 4 N ammonium hydroxide-ethanol eluate, which contained proctolin. The residue (approximately 30 g from the extract of 25000 cockroaches) was dis solved in 2 liters water and was applied to a 3 x 80 cm column of Dowex 50W-x8 (50 to 100 mesh, NH4+ form). The initial steps for the prepara tion of the resin for this column were the same as described above. The resin was then equilibrated with 2 N ammonium hydroxide and was washed thoroughly with water. After packing, the column was again washed with water. Following application of the sample, the column was eluted with 2 liters water and then with 0.05 N ammonium hydroxide at a flow rate of 75 ml/h. Fractions (25 ml) of the latter eluate were collected and bioassayed, and the eluate between 800 and 1600 ml that contained proctolin was combined. Removal of the solvent by using a rotary evaporator yielded about 700 mg residue. After use, the resin was recycled by washing consecutively with 3 vol. each 2 N sodium hydrox ide, water, 2 N hydrochloric acid, water, 2 N ammonium hydroxide, and water.
To process 125 000 insects (125 kg fresh weight) the above procedures were repeated four more times. By bioassay it was determined that the total residue (3.4 g) from five runs contained 1.3 x 106 rectum equiva lents, which is now known to equal 1.12 mg proctolin. At this stage the recovery was about 75% since it is estimated that one cockroach contains about 12 ng proctolin. In view of the recent paper by James (1978), it ap pears that a large portion of the loss of proctolin during this step may have
occurred as a result of the action on the arginyl moiety of ammonium hydroxide in the presence of the Dowex 50 resin.
b. Step 2. Chromatography over alumina was f~und to be an efficient way of further purifying the crude sample of proctolin. To avoid losses from autoxidation, 0.05% 4-methyl-2, 6-di-tert-butylphenol [frequently referred to as butylated hydroxy toluene (BHT», known to be useful as an antioxidant during chromatography of lipids (Wren and Szczepanowska 1964), was added to the methanol used for this step. A slurry of 300 g acidic alumina (80-200 mesh) activated at 150°C for 4 h to remove water was poured into a 3 X 40 cm column. The packed column was then washed with 500 ml methanol. The same solvent (500 ml) was added to the residue (3.4 g) obtained from the extract of 125000 insects as described in Step 1 and the insoluble portion (480 mg) was removed by fil tration by using a funnel with a fritted disc. After reduction in volume to 300 ml by rotatory evaporation, the solution was applied to the column. The column was first eluted with 400 ml methanol and then with a methanol-water gradient generated similarly to that described by Donald son et al. (1952). A constant volume of 3 liters was maintained in the mixer. The flow rate was approximately 125 ml/h and 25-ml fractions were collected by using a fraction collector over a 24-h period. By bioas say proctolin was found to be eluted between 1800 and 2400 ml, corres ponding to about 50% methanol. Removal ofthe solvent at reduced pres sure and room temperature with a rotary evaporator yielded 660 mg residue.
c. Step 3. The next stage of the proctolin purification employed a chromatographic grade cation exchange resin. Rexyn 101 resin (200-400 mesh) was cycled once as described for the Dowex 50 resin and then was converted to the NHt form by stirring with 2 N ammonium hydroxide. After the filtering and washing, a slurry of the resin in water was used to pack a 1.3 X 44 cm column. The active fraction that resulted from alumina chromatography was dissolved in 60 ml water and was applied to the column first eluted with 40 ml water and then with a water -0.04 N NH40H gradient over a period of 48 h at a flow rate of 12 ml/h. A con stant volume of 200 ml was maintained in the mixer and 9-ml fractions were collected by using a fraction collector. Solvent was removed in vacuo from that portion of the eluate between 270 and 400 ml that showed activity on the hindgut.
d. Step 4. Further purification was achieved by countercurrent dis tribution. This method depends on the partitioning of a mixture between two liquid phases and separations are obtained because of differences in the partition coefficients. The active residue (240 mg) from Step 3 was subjected to a total of 120 transfers by using 10 ml of each phase of the solvent system n-butanol-acetic acid-water (4: 1 :5, v/v) with a 60-tube automated instrument manufactured by H.O. Post. The use of such an
apparatus has been described in detail by King and Craig (1962). At the end of the distribution a smaIl volume was withdrawn from each tube, diluted with water, and assayed on the cockroach hindgut. Tubes 15-26 contained the major portion of the activity. The contents of these tubes were combined and the solvent was removed in vacuo by using a rotary evaporator.
e. Step 5. Paper chromatography was used for the next stage of the purification. Sheets of What man No.1 paper (15 x 35 cm) were washed with water and with 95% ethanol and then were dried at room tempera ture. Prior to the addition of the upper phase of the solvent system, n butanol-acetic acid-water (4: 1:5, v/v) containing 0.05% BHT, the large glass chromatographic tank was lined with Whatman 3 MM paper and flushed with nitrogen. The active fraction (35 mg) from countercurrent separation was dissolved in 1 ml 60% methanol containing 0.05% BHT and was applied as a narrow band to three sheets of the washed paper. These sheets were then placed in the tank and equilibrated for a 2-h period before development in the dark during a 16-h period was com menced. Examination of the developed chromatograms under u. v. light showed several zones. The area containing proctolin was located by bioassay in which very small pieces of paper removed from the chro matogram were placed directly in the organ bath. This zone (Rr 0.42-0.55) was cut out and extracted with water and the water was removed by lyophilization. When other areas of the chromatograms were sprayed with ninhydrin and heated in the oven at 110°C until maximum color de velopment had occurred, several colored zones were observed between Rr 0.11 and 0.42, indicating that paper chromatography had separated a number of inactive components. The ninhydrin solution used here and in subsequent steps was prepared by dissolving 0.3 g ninhydrin in 100 ml n-butanol and adding 3 ml acetic acid.
f Step 6. Further purification was achieved by high voltage paper electrophoresis, a technique used extensively in the isolation and iden tification of amino acids and peptides. For the work described here, a Savant Model HV 5000 TC high voltage electrophoresis system was used. Other similar equipment is also available commercially. Because of the danger inherent in using equipment operated at high voltage, it is im portant to closely observe precautions listed by the manufacturer in the instruction manual.
Sheets of What man 3MM paper (15 x 120 cm), prewashed with water, were used for the electrophoresis. The first separation was performed at pH 6.4 with pyridine-acetic acid-water (25: 1:350, v/v). The buffer was placed in the two chambers at the bottom of the tank and Varsol was layered over the buffer to a level sufficient to cover the cooling coils through which cold water circulated. The active lyophilized fraction from paper chromatography (5.3 mg) was dissolved in 1 ml 60% methanol con taining 0.05% BHT, and this solution was streaked with a Hamilton
syringe onto two sheets of paper about 15 cm from the anode end with allowance for the part to be immersed in the buffer. Amino acid references were applied to a separate sheet. The p~pers were then moist ened by spraying carefully so as not to disturb the origin line. After lightly blotting up excess buffer on each side of the origin line with another sheet of filter paper, the papers were placed immediately on the rack and set in the tank so that the top and bottom edges were immersed in the buffer. This resulted in a 100-cm distance between the anode and cathode. The electrophoresis was then run at 4000 V (70-75 rnA) for 2 h. At the end of the run the sheets were dried and the zone containing proctolin was located by bioassay as described above for the paper chromatograms. The reference compounds were located by ninhydrin spray; results are summarized in Table 1-2. Proctolin was eluted from the active zone with water. After lyophilization the active fraction was separated by high voltage paper electrophoresis at pH 3.5 with pyri dine-acetic acid-water (1: 10:445, v/v). In preparation for this step, the residue from the first electrophoretic separation was dissolved in 300 p.l 60% methanol containing 0.05% BHT and applied to a single sheet of Whatman 3MM paper. Standards were applied to a separate sheet. Both sheets were moistened with buffer by spraying and the electrophoresis was run at 5000 V (40 rnA) for 2 h. The proctolin zone and the position of the standards were located as before (Table 1-2). After extraction of the active zone with water, the quantity of proctolin present was determined by bioassay before lyophilization. Three other inactive zones well separated from proctolin were observed when the remainder of the sheet used for the proctolin separation was sprayed with ninhydrin.
Table 1-2. High-voltage paper elec trophoresis of proctolin and some amino acids.
Distance migrated toward cathode (cm)
Substance pH 6.4u pH 3.5b
Proctolin 19 32
Isoleucine 5 10.5
Histidine 30 52
Arginine 40 47
Lysine 43 50.5 aSolvent: pyridine-acetic acid-water (25: I :350, v/v). Electrophoresis was carried out at 4 kV(70-75 rnA) for 2 h. bSolvent: pyridine-acetic acid-water (1:10:445, v/v). Electrophoresis was carried out at 5 kV(40 rnA) for 2 h.
g. Step 7. The final steps were necessary mainly for the removal of contaminants apparently accumulated during electrophoresis. To avoid the introduction of further impurities it was necessary to ensure that only very pure solvents and chemicals were used and that all glassware was thoroughly cleaned. The presence of trace amounts of ninhydrin-positive impurities in the distilled water supply proved a difficulty. This problem has been discussed by Hamilton and Myoda (1974). Water for the remaining work was distilled after adding 0.25% solid sodium hydroxide and 0.05% potassium permanganate and then was redistilled twice with a glass system.
Gel filtration was useful for further purifying proctolin. Booklets available from Pharmacia, the manufacturer of Sephadex, provide a good introduction to the technique. A 1.6 x 190 cm column of Sephadex G 15 with a void volume of 160 ml was prepared and washed with 0.02 M am monium formate for 24 h. The ammonium formate used to prepare the eluant was freshly sublimed. Next, a 300-JLg sample of proctolin obtained in Step 6 was dissolved in 0.4 ml 0.02 M ammonium formate and applied to the column that was then eluted with the same salt solution at a flow rate of II ml/h. Fractions of 3 ml were collected. Small portions were removed for bioassay and the active portion of the eluate between 228 and 255 ml was lyophilized to yield 206 JLg proctolin (determined by bioassay).
With Sephadex G 15, elution volumes of I-JLmol quantities of (a) glycyl leucyl-tyrosine, (b) leucyl-tyrosine, and (c) tyrosine amide occurred in the expected order, a < b < c. The elution volume of proctolin was less than for these reference peptides, indicating that proctolin had a higher molecular weight than glycyl-Ieucyl-tyrosine (mol. wt. 351). Although this was subsequently shown to be correct, caution must be observed in attempting to estimate molecular weights by comparison of elution volumes of standards with that of an unknown substance unless all con tain an equal number of aromatic amino acid residues, since it is known that aromatic substances are reversibly adsorbed on Sephadex and thus are retained more than would be expected for their molecular weight.
h. Step 8. Remaining impurities were removed by passing the residue from Step 7 through a 0.2 x 20 cm column of Rexyn 101 (200-400 mesh, NHt form). A procedure similar to that described in Step 3 was used to prepare the column. After application of the sample containing proctolin, the column was washed overnight with 10 ml water. The column was then eluted during 24 h with a water-0.05 N ammonium hydroxide gradient at a flow rate of 0.35 ml/h. A constant volume of 10 ml was maintained in the mixer. Fractions of 25 drops were collected by using a fraction collector and were assayed on the isolated hindgut. Water was removed from the active eluate between 5.5 and 7.5 ml by lyophilization, yielding 180 JLg proctolin.
B. Characterization and Structure Elucidation of Proctolin
Some information about the chemical nature of proctolin was ac cumulated during the period required to purify this substance. For ex ample, it was recognized fairly early in the study that proctolin was a pep tide. However, it was not possible to make any attempt to determine the structure until a pure preparation had been obtained. Work that led to the structure H-Arg-Tyr-Leu-Pro-Thr-OH has been described briefly by Starratt and Brown (1975). This section presents additional details that, it is hoped, will be helpful to others trying to determine the structure of physiologically active insect peptides.
In work such as this when only a limited amount of material is avail able, methods must be chosen carefully. Also, since one cannot perform many trial experiments, all methods should initially be worked out with model substances. Finally, once a structure has been obtained that is con sistent with all the data and observations relating to the unknown, it should be confirmed by synthesis. Usually, one cannot be confident that the structure is correct until it has been shown that the physical, chemical, chromatographic, and biological properties of the synthetic substance are identical to those of the natural product.
1. Evaluation of Purity and Detection on Chromatograms
Several pieces of chromatographic evidence indicated that the isolated proctolin was sufficiently pure to permit an attempt to determine its struc ture. It gave a single ninhydrin-positive spot on high voltage paper elec trophoresis at pH 6.4 and 3.5 with systems described in Sect. III.A.2.f. Also, it was shown to be homogeneous by paper and thin-layer chroma tography. For the first of these latter methods, approximately 3 ILg proctolin were applied to sheets of What man No. I paper and chromato graphed with the upper phase of the solvent system n-butanol-acetic acid-water (4: 1 :5, v/v), as described in Sect. III.A.2.e. A spot for proctolin at Rc 0.46 was detected by bioassay and colored spots at the same position were obtained when the chromatograms were sprayed with either ninhydrin or the Sakaguchi reagent. Detection by bioassay and with ninhydrin has already been described. For the third means of detec tion, the thoroughly dried chromatogram was sprayed with a 0.1 % solu tion of 8-hydroxyquinoline in acetone and then, after drying, with a solu tion of 0.2 ml bromine dissolved in 100 ml 0.5 N sodium hydroxide. The positive reaction of proctolin to this spray, which detects unsubstituted or monosubstituted guanidines, was suggestive of the presence of arginine in this peptide.
About 3 ILg proctolin, as well as smaller amounts of several reference amino acids, were chromatographed on an A vicel thin-layer chromato graphic plate with n-butanol-acetic acid-water (4: 1: 1, v/v) and on a
Kieselgel plate with the upper phase of n-butanol-acetic acid-water (4: 1: 5, v/v). A ninhydrin-positive spot was observed at Rf 0.29 on Avicel and at Rf 0.17 on Kieselgel. Standard thin-layer chromatographic techniques such as those described in the useful handbook edited by Stahl (1969) were employed. Records of thin-layer chromatograms were usually made by one or more of three methods: (l) tracing the pattern of spots on trans parent paper, (2) photocopying, or (3) photographing. Although they may be obtained commercially, thin-layer plates used in this work were prepared with a Camag Automatic TLC Coater. Glass plates (20 x 20 cm) were coated with a 0.25-mm layer of Avicel (FMC Corporation) or Kieselgel DF-5 (Camag). The former were air dried and the latter were activated by being heated 1 h in an oven at 110°C before use.
Proctolin could also be detected on thin-layer chromatographic plates by bioassay. Small amounts of A vicel or Kieselgel were removed from the plate with a sharp spatula and were added directly to the bath contain ing the hindgut.
2. Ultraviolet Spectrum
The u. v. spectra of fractions that contain proctolin measured on a Gilford 2400 spectrophotometer showed an absorption peak at 277 nm suggestive of the presence of a tyrosyl residue. In the later stages of the large-scale purification of proctolin when only traces of impurities were present, chromatographic fractions could be monitored by use of an u.v. detector.
3. Hydrolysis of Proctolin
Most hydrolyses were carried out in 6 mm o.d. X 50 mm glass culture tubes (Kimax brand). Before being used these were cleaned with heat for 12 h at 550°C in a muffle furnace. Constant boiling hydrochloric acid was prepared by mixing concentrated hydrochloric acid with an equal volume of triply distilled water (Sect. III.A.2.g) and distilling under nitrogen (a slow stream of nitrogen was introduced into the distillation flask through a ground joint with a sealed inner tube that had been drawn out to a capillary). The fraction distilling at approximately lO8°C was collected and stored under nitrogen in a glass-stoppered flask. Dust was prevented from collecting around the top where it might fall into the flask upon open ing by a small sheet of plastic held in place by a rubber band.
For the determination of amino acid composition, 2-4 fJ-g proctolin in water were transferred to the small tubes and dried in the following fashion. After the aqueous solutions were frozen, the tubes were set in side an ice-cooled 25- or 50-ml conical flask and placed under vacuum (pressure less than 0.2 mm Hg) by attachment to the manifold of an Airlessware vacuum rack (Kontes Glass Co.). This system proved prac tical both for lyophilization and for evaporation of solvents in vacuo when working with a small number of samples. A dry ice-acetone bath
provided a fast and convenient means of freezing samples. When lyophilization was complete, 50 ILl constant boiling hydrochloric acid were added and the tubes were sealed in a fine oxygen-gas flame. Hydrolysis was accomplished by heating in an oven at 110°C for 16 h. The cooled tubes were opened by applying a hot glass rod (heated in an oxygen-gas flame) to a scratch made with a file or a diamond pencil near the top of the tubes and the hydrochloric acid was removed at room tem perature by using the vacuum system described above for the Iypohiliza tion. In a preliminary experiment, the residue from the hydrolysate was chromatographed on paper with the upper phase of n-butanol-acetic acid water (4: 1 :5, v/v). Ninhydrin spray gave several spots, indicating that proctolin was a peptide.
4. Identification of Amino Acids
The amino acids that constitute proctolin were identified by thin-layer chromatography of the dansyl (Dns) derivatives formed by reaction with dansyl chloride (l-dimethylaminonaphthalene-5-sulfonyl chloride). This simple and sensitive method has been widely used for the identification of amino acids (Rosmus and Deyl 1971; Niederwieser 1972). For this work a solution of 3 mg dansyl chloridelml acetone was prepared.
Dansylation was carried out essentially as described by Gray and Smith (1970). The residue from the hydrolysate of proctolin was dis solved in 15 ILl 0.2 M sodium bicarbonate and then dried in vacuo to remove traces of ammonia that might be present. The sample was redis solved in 15 ILl water, and the pH was checked by applying a small volume to short-range indicator paper to ensure that it was not below 7.5-8. Ifnecessary, the pH was adjusted to this level by the further addi tion of 0.2 M sodium bicarbonate. Then 15 ILl dansyl chloride solution was added to the sample with mixing by using a vortex mixer. The tube was covered with Parafilm and was heated on a magnetically stirred oil bath at 50°C for 15 min. A coil of copper wire wound so that the tube could not slip through and hung from the side of the oil bath was used to hold the tube. After the heating period the solvent was removed in vacuo at room temperature and the dansyl derivative was dissolved in ethanol water (3: I) and applied with a fine capillary to one comer of a 20 x 20 em silica gel 60 F-254 precoated glass plate (E. Merck) freshly activated at 110°C for 30 min. The plate was developed according to Gros and Labouesse (1969) with benzene-pyridine-acetic acid (80:20:5, v/v) and then in the second direction with toluene-2-chloroethanol-28% ammonia (6: 10:4, v Iv). After both the first and second developments the plates were dried in the fumehood by using a stream of air from a hair drier. Spots for the Dns-amino acids were located by examination of the plate under long wavelength u.v. light (366 nm) in a Model C-5 u.v. viewing cabinet (Brinkman). Comparison of the pattern to that for similar
chromatograms of 0.4 ILg Dns-amino acids, purchased from Nutritional Biochemical Corp. or prepared in this laboratory, and of material result ing from a control reaction carried out with only the reagents and solvents for hydrolysis and dansylation, indicated that hydrolysis of proctolin yielded arginine, leucine, proline, threonine, and tyrosine in approxi mately equimolar amounts. The chromatograph of the control reaction product showed the position of spots due to by-products such as Dns-OH and Dns-NH2 and demonstrated the state of purity of solvents and reagents.
To confirm the identification of the Dns-amino acids from proctolin, they were co-chromatographed with small quantities of the dansyl deriva tives of 14C-Iabeled amino acids, which were detected by au toradiography. Thus, L-[U-14C]-labeled arginine, leucine, proline, threonine and tyrosine (5-11 ng; 125-230 mCi/

neurohormonal techniques in insects

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