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
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© 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.
4 A.N. Starratt and R.W. Steele
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)
Proctolin: Bioassay, Isolation, and Structure 5
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.
6 A.N. Starratt and R.W. Steele
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-
Proctolin: Bioassay, Isolation, and Structure 7
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
8 A.N. Starratt and R.W. Steele
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
Proctolin: Bioassay, Isolation, and Structure 9
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
10 A.N. Starratt and R.W. Steele
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.
12 A.N. Starratt and R.W. Steele
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
Proctolin: Bioassay, Isolation, and Structure 13
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
Proctolin: Bioassay, Isolation, and Structure 15
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
16 A.N. Starratt and R.W. Steele
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
Proctolin: Bioassay, Isolation, and Structure 17
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
18 A.N. Starratt and R.W. Steele
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
Proctolin: Bioassay, Isolation, and Structure 19
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.
20 A.N. Starratt and R.W. Steele
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.
Proctolin: Bioassay, Isolation, and Structure 21
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
22 A.N. Starratt and R.W. Steele
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
Proctolin: Bioassay, Isolation, and Structure 23
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
24 A.N. Starratt and R.W. Steele
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/