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Advances in Insect Physiology

Volume 24

This Page Intentionally Left Blank

Advances in Insect Physiology edited by

P. D. EVANS Department of Zoology, The University Cambridge, England

Volume 24

ACADEMIC PRESS flarcourt Brace & Company, Publishers

London San Diego New York Sydney Toronto Tokyo

Boston

ACADEMIC PRESS LIMITED 24-28 Oval Road London NW1 7DX United States Edition published by ACADEMIC PRESS INC. San Diego, CA 92101

This book is printed on acid-free paper Copyright 0 1994 by ACADEMIC PRESS LIMITED

All Rights Reserved

No part of this book may be reproduced in any form by photostat, microfilm, or any other means, without written permission from the publishers A catalogue record for this book is available from the British Library ISBN 0-1 2-024224-9 Typeset by Columns Design & Production Services Ltd, Reading Printed and bound in Great Britain by TJ Press Ltd, Padstow, Cornwall

Contents Contributors Homologous Structures in the Nervous Systems of Anthropoda W. KUTSCH and 0. BREIDBACH

vi

1

Prostaglandins and Related Eicosanoids in Insects D. W. STANLEY-SAMUELSON

115

Cellular and Molecular Actions of Juvenile Hormone: General Considerations and Premetamorphic Actions L. M. RlDDlFORD

213

Mechanism of Action of Bacillus thuringiensis Insecticidal &Endotoxins B. H. KNOWLES

275

Insect Glutamate Receptors P. N. R. USHERWOOD

309

Subject Index

343

Contributors 0. Breidbach

Institut fur Angewandte Zoologie, Universitat Bonn, 53121 Bonn, Germany B. H. Knowles

Department of Zoology, University of Cambridge, Downing Street, Cambridge CB2 3EJ, UK W. Kutsch

Facultat fur Biologie, Universitat Konstanz, 78434 Konstanz, Germany L. M. Riddiford

Department of Zoology, University of Washington, Seattle, Washington 98195, USA D. W. Stanley-Samuelson

Department of Entomology, University of Nebraska, Lincoln NE 685834816, USA P. N. R. Usherwood

Department of Life Sciences, University of Nottingham, Nottingham NG7 2RD. UK

Homologous Structures in the Nervous Systems of Arthropoda W. Kutscha and 0. Breidbachb a

Fakultat fur Biologie, Universitat Konstanz, 7750 Konstanz, Germany lnstitut fur Angewandte Zoologie, Universitat Bonn, 5300 Bonn, Germany

1 Introduction 1 1.1 The phylum Arthropoda 1 1.2 New approaches, with a special consideration of the nervous system 2 1.3 The concept of the identified neurone 4 2 On the search for homology 5 2.1 From morphology to genetics 5 2.2 Development and immunohistochemistry 7 2.3 Definition of homology 10 3 Neural systems in Arthropoda 16 3.1 Insecta 16 3.2 Myriapoda 57 3.3 Crustacea 62 3.4 Chelicerata 69 3.5 Visual systems of Arthropoda 76 4 Conclusions 77 4.1 Segmentation 78 4.2 Homology 79 4.3 Phylogeny 80 Acknowledgements 83 References 83 Note added in proof 113

1

1.1

Introduction T H E PHYLUM A R T H R O P O D A

The Arthropoda constitute the phylum with the highest diversity of species in the animal kingdom (Boudreaux, 1979). Originated in the Precambrium (Bergstrom, 1979), this monophylum (Paulus, 1979; Ax, 1984) basically consists of two groups, the Chelicerata and the Mandibulata (Lauterbach, 1989). The common ancestor of the arthropods has to be regarded as an annelid-like species, with an exoskeleton, serial homologous segments ADVANCES IN INSECT PHYSIOLOGY VOL 24 ISBN W12424225-Y

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W. KUTSCH AND 0 . BREIDBACH

bearing a series of repetitively structured appendages, and possessing fused “head”-segments in the anterior region (Lauterbach, 1986). This ancestor must have displayed a specific type of photoreceptor comparable to that found in Onychophora (Eakin and Westfall, 1965; Paulus, 1979) and a ventral nerve cord (VNC) formed by a chain of serial homologous neuromers, the ganglia (Hanstrom, 1928a,b). Recent fossil evidence has debilitated an alternative view, the uniramian concept (Manton, 1977), which regards the Arthropoda as a polyphylum (Shear, 1992). According to the concept of an arthropod monophylum, compartments of the nervous system reveal some of the synapomorphies of this phylum (Hennig, 1950; Wiley, 1981; Ax, 1984). According to Hanstrom and later systematics, the common ancestor of the arthropods already possessed a brain, the principal organization of which was adapted in the different arthropod groups (Hanstrom, 1928a; Ax, 1984). The Onychophora represent a taxon which is regarded to have evolved convergently to the arthropod phylum (Boudreaux, 1979), new evidences even suggest that they are a sister-group of the Chelicerata (Ballard et al., 1992). Peripatus has a brain with several similarities to neuropile regions in the spider brain, for example the “mushroom bodies” and the “central body” (Schurmann, 1987).

1.2

NEW APPROACHES, WITH A SPECIAL CONSIDERATION OF THE NERVOUS SYSTEM

According to Hanstrom (1926, 1928a,b,c) such similar brain structures can be regarded as homologies throughout the arthropod species (Bullock and Horridge, 1965; Gupta, 1987b). Following the view of these anatomists, the common ancestor of the Arthropoda developed brain subcompartments, such as the central complex, the mushroom bodies and the optic lobes. However, recent analysis, based on a more thorough description of the morphology and the ontogeny of arthropod brains (for review, see Arbas er al., 1991), rendered such an interpretation as doubtful. When comparing the neuroembryology of different Mandibulata, one could establish some basic principles that might allow a homologization of certain cell lineages in this group. Thus, expression of homeobox genes like engrailed-originally described for Drosophila-were found to form equivalent patterns in the embryonic nervous systems of Crustacea, Myriapoda and lnsecta (DiNardo et al., 1985; Pate1 et al., 1989a; Whitington et al., 1991). Such new techniques in developmental biology provide a convenient addition to former methods of comparative neuroanatomy. In combining these new techniques with a morphological analysis, the fate of identified neural populations can be followed up in individuals as well as among different species. Thereby, based on comparative analysis, the development

HOMOLOGOUS STRUCTURES IN THE NERVOUS SYSTEMS OF ARTHROPODA

3

of the respective neural structures can be described. This review demonstrates that such a combination can prove to be highly successful in an identification of possible homologous structures in the arthropod nervous systems. Will such an extended comparative analysis provide enough material to understand neuroanatomical specifications of Chelicerata with respect to Mandibulata (Breidbach, 1992a)? Such a comparison can, however, become problematic. The morphological traits of Chelicerata development are entirely different to that of Mandibulata (Chabaud et al., 1990). Molecular biological techniques have never been employed to study the developing chelicerate nervous system. Thus, we do not know whether the DNA of species in that group encodes a set of developmental genes comparable to those of the Mandibulata. The morphogenetic features of neuroembryogenesis in spiders seem to be so peculiar that it appears unlikely that parallels can be found (Chaboud et al., 1990). Accordingly, the nervous system of the common ancestor of arthropods might have a much simpler structure than originally thought. If this is true, then such a view might implicate certain important aspects for a comparative analysis of the nervous systems of all these groups. The study of the nervous systems of species of the different Arthropoda groups may reveal certain variations of a restricted general theme, such as presented in the view that the “central complex” and “mushroom bodies” are homologous arthropod brain structures initially developed by a common ancestor. Alternatively, the different nervous systems might be the result of quite independent developmental strategies. If we adopt the latter view, then the structural equivalence in the organization of subcompartments that caused Hanstrom and his followers to develop their interpretations might reflect developmental or functional constraints in the organizations of the nervous systems (Hanstrom, 1928a; Bullock and Horridge, 1965; Gupta, 1987a). To decide how far these and other structural similarities in the morphology of the nervous system of different arthropods are due to homology, a comparative analysis has to be carried out and new technologies provided by developmental and molecular biology have to be included (Patterson, 1987). Identification of convergences in the structures of arthropod nervous systems enables one to describe the different ontogenetic and phylogenetic histories of these tissues. Based on such descriptions, we may be able to identify principles of common networks comprising the arthropod nervous systems. In addition, we might also be able to describe some of the functional constraints leading to apparently similar neuropile structures. If we can identify structurally similar neural structures that are the result of different ontogenetic programmes, then such analogous organizations may be associated with such functional constraints. We have to understand how far common characteristics of the performances of the arthropod nervous systems (Skinner, 1966; Paul, 1981; Meyrand and

W. KUTSCH AND 0. BREIDBACH

4

Moulins, 1988a,b) may occur with respect to different ontogenetic and phylogenetic strategies. 1.3

THE CONCEPT OF THE IDENTIFIEDNEURONE

An advantage of employing arthropods is that several of the relevant questions can be studied at the level of the identijied neurone (Wiersma, 1952; Hoyle, 1983). In its basic form, the concept of the identified neurone (Hoyle, 1983) tries to correlate the high degree of structural invariance of single identified neurones in one species with a functional specialization of this neurone. Consequently, a comparative analysis of neural architecture tries to reveal, to what extent differences in the construction of such identified neurones in different species can be regarded as the result of different functional specifications of these neurones (Dorsett, 1974; Weiss and Kupfermann, 1976; King and Valentino, 1983; Shaw and Meinertzhagen, 1986; Shaw and Moore, 1989). In this respect, a comparison of the organization of functionally different serially homologous neurones within one species becomes of interest. Using embryological and anatomical evidence, repetitively organized sets of neurones, with similar positions and fates, were identified in the different neuromeres of one individual (Taghert and Goodman, 1984; Pearson et al., 1985). Structural differences of such apparent serial homologues seem to be the product of functional specializations of the different segments and ganglia. This approach has been quite successful not only in a description of certain systems which trigger different behavioural programmes, but also in a description of the basic framework of an arthropod ganglion (Weevers, 1985). Furthermore, such an approach is advantageous, when combining the different results gained in the description of different functional units in different arthropod systems (Roeder, 1967). Thus, a basic scheme of structure and function of the nervous system in these invertebrates has been evolved (Edwards, 1977). The situation becomes different when we want to analyse the specificities of neural network organizations in different arthropods. Thereby, the complexity of the respective nervous systems has to be considered. Specificities may be the result of “internal” developmental constraints, following a developmental programme defined by the ontogenetic programme of an organism (Wake and Roth, 1989). This can be studied by an analysis of the different developmental histories of these species (Dohle, 1984; Dumont and Robertson, 1986), in which we have to deal with the radiation in the developmental strategies during the evolution of the Arthropoda (Anderson, 1973). Accordingly, it is not sufficient to search for common principles established in the organizations of the nervous systems of different arthropods. We, also, have to study the different life histories which,


5

eventually, might generate similar performances (Campbell and Hodos, 1970). In an approach to understand the pecularities of the different neural networks, such a comparative view is a prerequisite. Thus, we suggest that comparative neuroanatomy and physiology of the arthropod nervous system have to be paired with a thorough analysis of the different developmental and phylogenetic histories. This might result in an understanding of the constituents and the constraints which lead to the diversification of structures in the arthropod nervous systems (Arbas et al., 1991). Such a comparative analysis has to be based on phylogenetic systematics (Wiley, 1981). Apomorphies and synapomorphies or homologies (Hennig, 1950, 1982) have to be listed for the nervous system. If the variation of these structures is studied and if their fates throughout ontogeny are persued, then we may draw conclusions as to where and how far similar or divergent shapes of possible homologous structures reflect the different functional specifications of each individual species. In this regard, it seems most important to analyse the fate of such homologous structures for two reasons: (1) we may understand whether the radiation of species is reflected in the organization of their nervous systems; and (2) we may construct something like a framework of nervous structures that allows us to study in detail where, and at what stage of evolutionary development, new parts of the nervous system may have been added to the common “Bauplan”. The common “Bauplan” may be defined as the sum of all homologous structures in their original manifestation. Such an analysis can be extended stepwise from closely related species to larger taxons, studying the degree of variation found in the composition of apparent homologous structures. In such an attempt, the phylogenetically conserved basic structures will be characterized. These were varied in the different species due to adaptive radiation. This concept can give rise to new insights into the study of neural networks.

2

2.1

On the search for homology FROM MORPHOLOGY TO GENETICS

How does one start such an analysis? One has first to find reliable criteria to compare the organization of nervous systems in different arthropods. These criteria will only be valid if one can identify homologous structures in the different nervous systems. Ideally, these homologous structures should be homologous neurones, whose fate can be followed up and whose divergence in different groups can be studied in detail (Croll, 1987). The arthropods provide a unique opportunity to describe whole sets of such possibly homologous neurones (see Bullock, 1978; Northcutt, 1984). Identified neurones can be described by morphological and developmental criteria (Croll, 1987). Thereby it becomes possible, not only to describe

6

W. KUTSCH AND 0.BREIDBACH

serial homologous neurones within one species, but also to depict homologous neurones in different arthropod groups. Hence, the combination of new techniques, as provided by modern immunohistochemistry , neuroembryology and molecular biology, combined with morphological analysis can give new and reliable tools to identify such homologous structures in the invertebrate central nervous system (CNS) (Weiss and Kupfermann, 1976). Neuroembryology allows the fate mapping of certain neural clones (Stent and Weisblat, 1985). It is possible to follow the fate of identified neurones and to describe the growth of their branching patterns (Goodman and Spitzer, 1979; Bate et al., 1981; Goodman, 1982; Shankland and Goodman, 1982; Taghert and Goodman, 1984). Such studies, carried out in different insect and crustacean species, show the developmental patterns of the Mandibulata nervous system with a strict stereotype (Thomas et al., 1984). The timing of development of certain types of neurones is accurate and occurs at a specific time scale in the different ganglia of an individual (Bastiani et al., 1985). Thus, the neural architecture of the imaginal nervous systems shows not only a repetitive structure, but the reiterated components are found to be based on the common developmental programmes of the serial homologous neurones. It is obvious that the pattern of neural connections is much simpler in the ganglia of the ventral nerve cord (VNC) compared with the highly diversified arthropod brain. A comparison of VNC units with brain subunits might offer the possibility of exposing the underlying principal segmental nature of the arthropod brain neuromeres. This would contribute information on the questions of the principal constituents of the arthropod head (Schmitt-Ott and Technau, 1992). This may also have some impact on the analysis of the phylogenesis of the arthropod species (Weygoldt, 1979). In addition, such an analyis may provide the basis for a detailed comparative analysis of arthropod brains (It0 and Hotta, 1992). Yet, what are the criteria for the identification of homologous structures in nervous systems? A principal advantage of employing arthropod species is the possibility of studying identified neurones. Thus, physiologists work on defined elements of a neural network to reveal the functional architecture of its basic units. A former concept, in which the precise pattern of identified neurones was deduced from a functional specification of these neurones (Kupfermann and Weiss, 1978), has had to be modified recently, since serially homologous neurones have been identified that show structural similarities in spite of the functional diversification of the appropriate ganglia (Wilson and Hoyle, 1978). Goodman (1978) and Macagno et al. (1973) demonstrated the degree of variability in the architecture of identified neurones in clones of insect or crustacean species. Structural uniformity of identified neurones, even within one species, seems to be restricted to certain basic patterns governing principal pathways of axonal projections


7

and the general dendritic arborization pattern (Satterlie, 1985). Accordingly, the functional peculiarities of one neurone appears to be primarily due to its contacts within the neuropile (Pearson and Goodman, 1979). Therefore, further analysis of the functional organization of arthropod ganglia should be centred on physiological descriptions of neural connectivity (Ewert , 1980; Camhi, 1984).

2.2

DEVELOPMENT AND IMMUNOHISTOCHEMISTRY

Developmental biology and immunohistochemistry have introduced some new aspects in our understanding of structural specifications of identified neurones. It became possible to describe peculiarities of the arthropod neuropile in biochemical terms (with respect to the abundance of neurotransmitters and neuromodulators, see Rowell, 1976; Bicker and Menzel, 1989). Sets of repetitively organized neurones were found sharing structural and biochemical features. Supplemented by analysis of the developmental fate of several neurones, serial homologues can be shown, at least for the VNC (Nbsel, 1987b; Breidbach, 1990a). Recently, it was attempted to extend such an identification of serial homologous neurones in the VNC to include brain interneurones. Thereby, the principal neuromeral constitution of the brain ganglia may be elucidated (Breidbach, 1990b). These attempts were substantiated by techniques employing developmental genetics (Schmidt-Ott and Technau, 1992). Serial homologies originate by repetitive, equivalent developmental programmes in the different segmental units of the body plan of a species (Taghert and Goodman, 1984). Thus, one can make inferences on the degree of morphological variance found in a population of ontogenetically uniform cells based on a genetically uniform organism. Furthermore, studies of serial homologies result in the establishment of morphological criteria for a description of subcompartments in a tissue which can vary morphologically on a larger scale (Breidbach and Dircksen, 1991). Serially homologous neurones are expressed in fused and unfused ganglia, thus allowing a description of their correspondence in different neuropile segments. Thereby, we can gain additional criteria to identify subunits in an otherwise extensively varying tissue (e.g. development of the sub- and supraoesophageal ganglion). As synapomorphic attributes of the nervous system of one group of arthropods can be described by developmental analysis, trends in the neuronal organizations of the related species can be viewed as being due to functional adaptations. We know, for example, that the principal structural and biochemical constitutions of interneurones (INS) of insects and crustaceans as well as those of insect larval motoneurones (MNs) are established in the late embryo, and are maintained throughout ongoing

8

W. KUTSCH A N D 0.BREIDBACH

development (Breidbach, 1990a). During metamorphosis of holometabolan insects, these neurones persist and extend their dendritic fields (Levine, 1986, 1989; Breidbach, 1987a, 1990~).Accordingly, a description of the larval or the late embryonic organism outlines the principal shape of such identified neurones (Breidbach, 1991). For our approach, this is of special importance since the larval and the adult instar might have been subjected to different functional constraints. Such different histories might have resulted in morphologically and functionally highly different types of organisms throughout ontogenesis, such as is apparent in the caterpillar and the butterfly. In systematics, Hennig (1982) therefore proposed the semaphore concept, thereby warning that comparative morphological analysis may mix characteristics of different ontogenetic stages in a cladistic analysis. Neuroanatomists have to be aware of this problem, too. Here, we argue that the late embryo or the first larval instar represent the basic type of the nervous system of an arthropod. Later transformations only seem to vary this principal structure (Breidbach, 1990~).They may be due to shifts in the sensory inputs, as performed in the holometabolan insects (eye and antenna). Thus, for a comparative analysis of the neuroanatomy of different species, it seems obvious to concentrate on comparisons of the late embryo or first larval stage. Fate mappings of identified neurones can be combined to describe the complete history of certain neural populations. By a combination of single cell labelling techniques with histological methods, the life history of certain neurones can be followed from their birth onwards (Stent and Weisblat, 1985). It becomes possible to identify the progeny of certain neuroblasts and, thus, describe the lineage to which certain neurones belong (Truman, 1990). A comparison of the cell lineages in different species establishes the strongest criteria for comparative phylogeny (Thomas et al., 1984). In Mandibulata, the early patterning of neurogenesis appears to be quite similar on the morphological and gene expression levels (Campos-Ortega et al., 1989). Genes regulating neural development, such as engrailed or wingless, are expressed in a wide range of different Mandibulata species, for example Crustacea, different insects (Locusta, Tribolium, Diptera) and myriapods, with a similar timing of differentiation (Patel et al., 1989a; Whitington, 1991; Stuart et al., 1991). Therefore, development of their nervous systems seems to be established by common mechanisms. Comparative genomic analysis establishes that, at least within insects, similar antigen expression patterns are based on evolutionary highly conserved DNA sequences (Patel et al., 1989a; Stuart et al., 1991). With such segmental markers coordinates are represented, by which neuroblasts could be homologized using positional criteria for a comparison of neuronal maps. Accordingly, homologous neuroblasts may be identifiable in different arthropods. Consequently, the progeny of such homologous neuroblasts may also be regarded as homologues.


9

Therefore, the criterion for the identification of homologous neurones is a common ontogenetic origin. How can we identify such homologous neurones in different species? Cell lineages can only be determined when criteria are available to define common coordinates for the early nervous systems of different species. Later, correlation becomes possible by the identification of neuromere compartments in the different species. Since these compartments are correlated, references are available to identify homologous neuroblasts and their progeny. The criteria to identify such homologous structures in the nervous systems of arthropods are basically morphological. The map of neuromeral subcompartments must be described. The scheme for such a map may be obtained by the analysis of expression patterns of developmental genes, such as engrailed. However, a restriction to the expression of a single gene would be insufficient. This would imply that genes like engrailed are always expressed by cell populations sharing the same relative position in all the different embryos (Baker, 1991). However, this need not be true in all cases. A gene, such as engrailed, determines the ongoing development and thus establishes some coordinates (Levine and Macagno, 1990), but the specificity of such gene expression may vary in different arthropods (Whitington et al., 1991). Thus, we have to find additional criteria for ganglionic coordinates. We have to find markers that are expressed in later stages of development, when there is no expression of the developmental genes like engrailed or wingless. A multitude of such criteria is not only useful because of practical reasons-but also in a more general perspective. The parallel structuring of the arthropod neuromeres leads to the formation of a repetitive neural organization as expressed by the serial homologous neurones. The serially homologous neurones are effective markers even in the adult nervous tissue and allow the application of the principal scheme of neuropil organization in the differentiated non-embryonic nervous systems. By their highly diversified morphologies, these serial homologues provide a fine scaling for a comparative analysis of arthropod nervous systems throughout the whole of postembyronic development. How is it possible to identify such serially homologous neurones? We can study motoneurones that are identified by the position of their somata, the pathways of their axons, the areas covered by their dendritic projections and by the topologies of their peripheral targets. Thus, for Crustacea (Miller et al., 1985) and insects, serial homologues of motoneurones innervating the dorsal or ventral longitudinal muscles can be determined (Davis, 1983; Breidbach and Kutsch, 1990). Furthermore, we can analyse interneurones characterized by common neurotransmitters. Th&e neurones can be identified by immunohistochemistry, in addition to the topological criteria. Thus, evidence is provided for homologous neurones, initially for closely related species. This will result in a description of a network of serially homologous neurones sharing characteristics of the network described

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originally. Accordingly, such an analysis can be extended to include more distantly related species. Such a comparative neuroanatomical approach searching for criteria of homology will include the identification of common neuroblast progeny. During neurogenesis of myriapods and insects, adaptations of the embryonic development may cause specific variations in the time course determining the early structural patterning, differences occurring even in closely related groups; see, for example, the development of contralateral projecting neurones in different myriapod species (Korschelt, 1936). Different structures in the adult nervous system are due to alterations of the ontogenetic programme. However, a purely descriptive embryology may have difficulties in untangling whether different courses of development are in fact due to principally different decisions in early neurogenesis. Thus, it is doubtful whether different morphogenetic features found in the ontogenesis of myriapod species are due to different programmes of the interacting nerve cells or are just reactions to the presence or absence of huge bulks of yolk, the extensions of which may vary considerably in different myriapod species. A comparative analysis of gene expression, using for example immunohistochemistry, does not provide conclusive evidence of homology (Baker, 1991). The failure of any immunopositive signal might be due to the absence of the reaction, or to slight variations in the structure of the respective proteins that inhibit antibody labelling. Alternatively, it could be due to methodical problems (fixation, buffers, etc.; Baker, 1991). Additional criteria, such as neuroanatomical descriptions, are necessary to evaluate the viability of the developmental approach in each case. If properly carried out, either approach-the embryological or the morphological-may be sufficient on its own, at least at the level of closely-related animals. However, a combination of both approaches is necessary when the comparison is carried out between higher levels of taxa.

2.3

DEFINITION OF HOMOLOGY

2.3.1 Historical aspects What is the meaning of homology in this context? The history and the lasting discussion of this concept in morphology has been reviewed repeatedly (Campbell and Hodos, 1970; Patterson, 1982, 1988; Croll, 1987; Wagner, 1989). They all refer to the classical definiton of Richard Owen (1843), who defined a homologue as “the same organ in different animals under every variety of form and function”. The historical presentation of a non-evolutionary morphology has been discussed by Patterson (1988). This concept of homology is based on the idea of a type (Typus) of organism, outlined in the famous academic dispute between Geoffroy de Saint Hilaire


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and Cuvier, 1833, in Paris (see Breidbach, 1982). The problem of a proper description of the development of the homology-concept occurs since this concept was established before the idea of evolution was introduced into morphology (Haeckel, 1866). Accordingly, the criteria used by previous authors often do not explicitly refer to evolutionary terms. This arouses some obscurity especially in dealing with convergence (Wagner, 1989). Recently, these problems gained some new support, since molecular analysis has succeeded in revealing systematic relationships using numerical methods (Reeck et al., 1987; Dover, 1987). At a first glance, systematic distances seem to be measurable by quantitative means (Smith et al., 1981). However, these ideas were falsified (McKenna, 1987; Sibley and Ahlquist, 1987). It has now become clear that the conceptual background of comparative analysis has to be based on a morphological methodology (Patterson, 1987). Thus, while molecular methods can provide new and detailed data that may fit into the respective conceptual frameworks, it is clear that the data obtained from older methodologies cannot be discarded. In the present chapter, we define homologies in a phylogenetic, systematic sense, representing synapomorphies (Hennig, 1950, 1982). Thus, we use the term homology to indicate a fundamental similarity due to inheritance from a common ancestral form. Theroretically, it might be possible to prove homology even for individual neurones, if their development is solely governed by specific genes. This appears to be the case for a subset of neurones in Drosophila melanogaster, whose fate is controlled by the segmentation gene even-skipped (Doe et al., 1988). We can study the development of certain neurones and describe their structure as the result of a gene cascade. The occurrence, specificity and sequence identity of the genes in this cascade can be studied, but this will only give some indication of whether or not certain neurones are homologous in different species. We need to extend the criteria for homologies provided by a similarity in details and in unique features (Remane, 1956; Simpson, 1967). These morphological features can only be regarded as the product of a specific ontogenetic programme, depending on gene expression and modified by cell-cell interaction (Breidbach, 1990a). Evolution of morphologies has to be understood in the light of the evolution of ontogenies. Accordingly, we should compare the developmental programmes of the neurones we suppose to represent homologues. Thus, homologous neurones are defined as sharing a similar development and originating from homologous precursor cells. Neuroanatomy provides only a method to uncover possible homologies in different species and we have extended the definition of this formerly exclusive morphological term. As discussed by Wagner (1989), some difficulties are implicit when adapting the homology concept to a strictly evolutionary perspective. However, since we identify homologous neurones

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by morphological criteria, these difficulties in our approach are not of methodological origin. Homologies depend on ontogenetic strategies that reflect common phylogenetic origin. If throughout evolution ontogenetic strategies are varied, this will cause different imaginal morphologies, which can be detected by a comparative embryological approach. The concept of DNA-sequence homologies does not fit completely into such a picture. DNA similarities-isologies in the sense of Wegnez (1987)-0nly indicate equivalent developmental strategies. They do not allow an understanding of the instant developmental pattern, within which these similar genes are expressed (Baker, 1991). DNA-hybridization, DNA finger printing and other methods have become established as new tools. They may become very effective for the description of complete DNA expression patterns (for review, see Patterson, 1987). These tools may establish a very sensitive addition to the recent morphological methodology, but even these tools cannot change the principal morphological character of the homology concept (Patterson, 1988). As mentioned above, homologies are defined as synapomorphies (Hennigian concept, 1982). Previous attempts to homologize neuropile structures on gross morphological criteria, such as determining the distribution of aminergic neurotransmitters in Crustacea (Aramant and Elofsson, 1976a,b; Elofsson, 1982) or the histological gross organization of neuropile structures in Arachnida or Mandibulata (Holmgren, 1916; Hanstrom, 1928a; Cupta, 1987b) appear to be vague, since the resolution of these methods is not accurate enough to provide detailed information about the cellular organization of the respective elements. With the possibility of identifying individual cells and describing their fate starting from the respective neuroblasts, a comparative analysis has gained new and unforeseen detailed material to reveal the radiation of the arthropod “Bauplan” of the nervous system. To understand the intrinsic capacities of neural networks, it is essential to establish the particulars of the appropriate neural “Bauplan”. 2.3.2 Criteria for homology For a description of structures within different species it is often of paramount interest to understand whether these structures can be compared in more detail. There are several aspects which might be of importance: anatomy-histology, function, and ontogeny. Inevitably, such comparison will lead to questions concerning the status of possible homology. Homologous structures sensu strictu are only those that descend from a common precursor cell or tissue (e.g. Arbas et al., 1991). Such a definition will allow a list of “ontogenetic homologues” when following up the development of an individual. However, evidently, such a definition must exclude a comparison among species. Comparison between different species


13

is not sufficient to describe the general “Bauplan” of a species. The definition of homology provided by Owen (1843: “the same organ in different animals under every variety of form and function”) could not solve the problem since this other definition merely replaces ‘‘homologous’’ by “same”. Owen (1848) himself tried to overcome this problem by applying two criteria towards “sameness”: relative position and connectivity (see also Boyden, 1969; Croll, 1987). Ultimately, with respect to phylogeny the term “common precursor” cannot be restricted to an individual’s life but would mean a common ancestor from which homologous structures develop from genomically identical sequences (Dumont and Robertson, 1986). Eventually, it should become possible by comparison of different related animals to search for a more general “Bauplan”, the basic organization of a common ancestor (Wagner, 1989). It would be of great advantage if studies of homology could always be paired with structural correspondence of proteins (Winter et al., 1968; Margoliash, 1969; Patterson, 1987; note, however, the controversy: Aboitiz, 1987; Dover, 1987; Wegnez, 1987; and the problems of protein sequences compared with evolutionary events: Doolittle, 1981). However, as has been discussed, since such an approach is not carried out routinely, we have to rely on morphological (occasionally aided by physiological) criteria to claim for homologous structures. “Homologous similarities are inferred inherited similarities that define subjects of organisms at some hierarchical level within a universal set of organisms” (Eldredge and Cracraft, 1980). However, which method can be used to distinguish homology from other similarity (analogy, homoiology, homoplasy: similarity of structures due to common functions, independent of inheritance; Simpson, 1967; Ax, 1984)? Simpson (1967; see also Dobzhansky et al., 1977) mentions two criteria which may help in a decision for possible homology: (1) with respect to any given feature minuteness of resemblance suggests homology; and (2) the greater the number of features (multiplicity of similarities) between any two organisms, the more likely it is that homology, caused by common ancestry, is responsible for the similarities of any one feature. It is quite obvious that we have to deal with empirical data which are often based on probabilistic inferences; for the concept of cladism, see Ax (1984). Remane (1956) compiled a catalogue of three criteria to define possible homologies: 1. Criterion of position: homology may be inferred for a structure in an equivalent position within a comparable framework. 2. Criterion of specific quality: similar structures may be homologized when they match in several specific features, irrespective of position within the organism. Trustworthiness increases in parallel to the grade of complexity and convergence of the compared structures. 3. Criterion of continuity: dissimilar and differently placed structures may be homologous as long as there are intermediate stages satisfying one or both of the aforementioned criteria. Such intermediate stages may

14


be due to ontogenetic processes or found as genuine systematic interjacent stages. Besides interspecific homology, serial homology should also be mentioned. Serial homology comes into play in segmented animals. It describes the relation between repeated structures, such as muscles or appendages along the segments of the Arthropoda or the limbs of Vertebrata. They obviously do not stem from the same precursor but equivalent precursors have to be assumed. Here, too, “equivalent” neurons can be identified only in a comparative developmental analysis. Nevertheless, in the following pages many cases will be presented which are regarded as serial homologues accepting a situation in which, during ontogeny, equivalent (=homologous) precursors have to be assumed for the different segments. 2.3.3 A catalogue for homology in terms of neural structures

In view of the criteria of homology outlined above, it is quite obvious that it will often be difficult to claim homology within the nervous system (Campbell and Hodos, 1970). In this case specific considerations are required since, unlike more compact structures such as bones, muscles or organs of vegetative functions, the nervous system is composed of large numbers of cells that interact via different types of processes. There are short dendritic processes, and the much longer axons which may connect neurones over long distances, reach the peripheral musculature or represent the afferent fibres from distal fields of sensory receptors. Additionally, especially in the group of Arthropoda, it often happens that parts of the CNS contract during ontogeny. Individual ganglia may fuse with each other. For example, in Locusta the first three abdominal ganglia fuse with the true metathoracic neuromere. In spiders and Diptera the CNS becomes an almost compact mass of neuronal and glial cells. Nevertheless, it is possible to attempt to identify homology on certain criteria. Generally, the criterion of position can be used, especially with respect to an existing frame work. More specifically, the following aspects need to be considered: 1. The position of a certain cell body may be employed in relation to other somata, to ganglionic structures (nerve roots, commissures, connectives, fibre tracts, penetrating tracheae). 2. The size of a soma, also with respect to its surroundings, may provide some hints. 3. The number of major dendrites, their positions along the neuropilar segment , the characteristics of neuropile penetration and the structure

of the complete dendritic field may be of importance.

4. The couse of the appropfiate axon(s) may have some meaning with respect to the ganglionic framework (for example, for INS a scheme has been offered; Robertson and Pearson, 1983). For MNs, the fact that they exit via a specific nerve root can be used as a rough


15

classification. Several axons may run close together as a group and each axon may be identified from its position within the group and its relative size. 5 . In the case of MNs the pathway of the axon in the peripheral nerves as well as the termination of the axons along the different muscles can be referred to. With respect to ontogenetic early interactions among cells and tissues it is to be expected that homologous muscles will be innervated by homologous MNs (see also Paul, 1991). For muscles, it might be easier to claim homology based on topographical as well as functional correspondence (Mickoleit, 1961). Actually, in terms of serial homology, muscles were among the first examples used for comparative and functional studies (for Orthoptera: Voss, 1905; Snodgrass, 1929, 1935; Albrecht, 1953). Therefore, by considering adjacent muscle homologues which also require ontogenetic early stages, identified homologous MNs may be discovered. 6. Additionally, physiological criteria of a neurone may be evaluated. MNs may be characterized as fast or slow excitatory or as inhibitory neurones (Hoyle, 1957). Other possibilities of discrimination would be spiking vs. non-spiking neurones (Burrows, 1985). Neurones may be involved in distinct motor patterns, whether as “command neurone” or elements of a neuronal circuit, and this may aid in a search for possible homologous (including serial homologous) neurones. 7. It is obvious that neurones can contain different transmitters (Nassel, 1987a). Additionally, it becomes evident that colocalization of neuromodulators and neurotransmitters occurs (Homberg et al., 1990). Such biogenic substances even were thought to indicate the progeny of a single neuroblast; however, recent studies demonstrate that such a rigid assumption has to be discarded, since the transmitter-type within such a family may well be different (Stevenson et al., 1992). Nevertheless, a coincidence of several criteria-morphological, immunohistochemical and possibly physiological-may strengthen a case for potential homologues. 8. Recent techniques in molecular biology open new fields with respect to homology studies. The expression of specific genes during specific phases of ontogeny might point to a distinct class of neurones, that may even indicate individual neurones (for instance even skipped). Such markers, like engruiled, have already been used to demonstrate parallel processes in a segmented insect (serial homology in the locust) but also are seen in crayfish (Pate1 et al., 1989b). The opportunity to mark specific individual neurones throughout early neurogenesis (Meier and Reichert, 1991a) opens up the possibility to compare different species in the search for such individual cells. The above list demonstrates the possibilities that exist, from a histological description to genetic approaches, to distinguish between individual

16


neurones. There is a great advantage in the study of the invertebrate nervous system compared to the vertebrate nervous system: in the invertebrates the total number of neurones is smaller. This makes it much easier to study the nervous system at the level of small neuronal networks. From here, it becomes possible to study identified neurones; and this must ultimately be the basis on which comparative studies can be used to identify homologies within an animal, as well as among different ones. As mentioned above, similarity, based on structural features, is one of the main arguments for possible homologues (although dissimilarity does not exclude it; Smith, 1967). This raises the question of how far can deviation of single elements be tolerated before one can discard an assumption of similarity? The natural variability must be studied by comparing identified units of a single species. Thus, an indication of endogeneous variation will be given which may help to decide whether serial or interspecific candidates can be considered to be homologous. By employing isogenic grasshoppers, clutches with duplications or deletions of large ocellar INS have been observed, suggesting some genetic control of a selective additional cell division or selective cell death (Goodman, 1974, 1977). Other work shows that the main branches of a neurone are rather invariant, while some of their side branches may be more variable or even missing (Goodman et af., 1979; Steeves and Pearson, 1983); with respect to a wild population, this variability appears to be even reduced (Satterlie, 1985). These examples demonstrate, that variation of identified elements may occur, albeit restricted to side branches which may have no major impact on motor control (Steeves and Pearson, 1983). For the wing stretch receptor, the constancy of central branching is striking, radical departures from the normal pattern (“mistakes”) are rare (c.3%) (Altman and Tyrer, 1977a,b). Therefore, to look in the adult instar for possible homologues does not require an absolute matching but a matching of some key figures. These, probably, can best be described by reverting to embryogenesis when the basic circuitry is laid down. This method of studying early neurogenesis may also help to decide on possible homology even in cases where further development will result in different morphology and synaptic connections (Pearson et af., 1985).

3 Neural systems in Arthropoda 3.1 3.1.1

lNSECTA

General aspects

There is a large amount of literature in which the nervous system of Insecta has been described, from the primary apterygote Lepisma (Barlet, 1951,


17

1953, 1954) to the most advanced Diptera (Power, 1948). A review on the older literature is given in the relevant chapters in Bullock and Horridge (1965). Such descriptions provide a framework for studies of the appropriate peripheral targets (sensory, muscular or endocrine system). Especially with respect to the thoracic nerves, similarities can be seen when comparing different insects, pointing towards homologies (Markl, 1966). There are “reasonable homologies on the basis of similar or nearby muscle block and sensory area but exact homologies are seldom certain because the distribution of the branches of a nerve in one genus is rarely identical to that in another” (Bullock and Horridge, 1965, chapter 16, p. 870). It is of interest that as a major argument for possible neural homologies, the relation of a nerve with a muscle group, or sensory area, is quoted also as well as the pattern of nerve branching. However, with respect to a more sound argument on homology, it is necessary to reach the level of identified central or peripheral (=sensory) neurones. The total number of neurones in Insecta is still vague. Generally it is accepted that the number reaches c.106 cells, with the largest part being found in both fused ganglia of the head (supraoesophageal = brain and suboesophageal ganglion). For the thoracic nervous system of locusts 1000-1500 neurones have been determined for a hemiganglion (Bate et a f . , 1981; Thomas et al., 1984), while a non-fused hemiganglion of the abdominal nerve cord contains only about 250 neurones. For the holometabolous insect Manduca, Truman and Booker (1986) estimate that the first thoracic hemiganglion contains 750 neurones in the last larval stage, and 2500 neurones in the newly emerged adult (a similar number is given for young adults of Drosophifu;Prokop and Technau, 1991). The latter increase is due to the renewed activity of neuroblasts. However, the total number in adults may decrease due to cell death of newly born neurons, such as is observed for a Manduca abdominal hemiganglion: compare 450 neurones at the pupal stage, and 250 neurones in the later adult stage, respectively (Truman, 1983). Therefore, the total number of VNC neurones might not be very different, when comparing Hemi- with Holometabola. 3.1.2 Motoneurones Staining methods (such as Timm’s intensification (Timm, 1958) of a cobaltor nickel-stained preparation; see Quicke and Brace (1979) in combination with Bacon and Altman, 1977) can be employed to count the number of MNs. The total number of MNs for a thoracic segment is in the range of 130-150, while for the abdomen there are about 70 (Schistocerca, Kutsch, unpublished observation). A comparison of existing data shows considerable differences in estimates, even from the locust which may be partly due to different staining techniques. A further problem arises from the minuteness of several fibres which may often result in incomplete central stainings in

18


backfills. In addition, too intensive central staining often prohibits a clear visualization of individual somata. Occasionally, one might have to deal with the problem of transsynaptic or interneural migration of the metal ions (Strausfeld and Obermayer, 1976). Exact cell counts may also be impaired by the fact that some neurones (such as DUM or VUM, see below) may innervate several muscles exiting via different nerve roots (Watson, 1984; Pfluger and Watson, 1988; Ferber and Pfluger, 1990; for cases of most complex peripheral organization, see Braunig, 1988, 1990). 3.1.2.1 Serial homologies There is a relatively large amount of knowledge suggesting apparently homologous muscles may represent serial homologies. A long search was made for the corresponding parts of organisms “which are built according to the same body plan” (Wagner, 1989). In segmented animals such a “body plan” could be, the “segmental plan”. Therefore, comparative studies of the musculature, their innervation and the associated MNs could result in a demonstration of a groundplan of an insect (or any other segmented animal) (Wittig, 1955; Bullock and Horridge, 1965; Markl, 1966). However, it becomes apparent that this wealth of information on the peripheral side is not paralleled by studies of the CNS. Information on serial homologous MNs is scarce. Therefore, it is still not yet clear whether there is a basic neural organization valid for all segmental ganglia. With respect to the CNS of locusts, it was shown that during neurogenesis each segmental ganglion provides a more or less identical set of neuroblasts, which is composed of 30 neuroblasts per hemisegment and an additional median neuroblast (Bate, 1976; Doe and Goodman, 1985; Shepherd and Bate, 1990). As mentioned above, the final number of neurones in the thoracic ganglia is different by a factor of about 4-5:l when compared to the abdominal ganglia. Several processes are known which may explain such differences in neural numbers starting off from an almost equal neuroblast array (Booker and Truman, 1987; Shepherd and Bate, 1990; Truman, 1990): 1. Throughout the animal kingdom the process of cell death is ubiquitous and this also occurs in the nervous system. During neurogenesis an overproduction of neurones may occur, a large number of which may die sooner or later (for some possible reasons of neuronal death, see Truman, 1983; Kutsch and Bentley, 1987). 2. Also, other processes, such as differences in the life time of individual neuroblasts and their division rates may result in segmental differences in final neurone numbers (Shepherd and Bate, 1990). Is there really a basic “Bauplan” of the segmental nervous system? Irrespective of the method used to reach a different number of neurones for the different segments, it needs to be asked whether the concept of serial homology for each individual neurone can be accepted. If so, then the divergence between the segments needs to be explained. Three possibilities should be considered:


19

1. The original structure valid for all ganglia is represented by each abdominal ganglion. In the thoracic ganglia many more neurones are added to enable various specific functions such as flight, walking, etc. to be carried out. 2. The neuronal organization of the thoracic ganglia is typical for all ganglia. Supernumerary neurones are lost in the abdomen during development, according to any of the cases of cell death, mentioned above. 3. Depending on specific genetic programmes, neuroblasts and neurones are committed to different fates in the different segments. This, consequently, would lead to the knowledge that for a large percentage of the neurones no homologues exist at all, for the different segments. Counts of the MNs indicate (see above) that the difference between thoracic and abdominal ganglia is not so evident, as compared with the total number of neurones. However, there is still a factor of 2-3 in which the thoracic system surmounts that of the abdomen. There are only a few cases described in which selective stains of the nerve branches reach individual muscles resulting in the claim for homologous MNs. 3.1.2.1.1 Typical rnotoneurons (MNs). On morphological criteria, Simmons (1977) described several of the MNs supplying the major flight muscles in dragonflies. He showed that the number of cell bodies innervating homologous muscles (of the meso- and metathorax) as well as their position were very similar. The dendritic organization of these MNs has not yet been elucidated so clearly, though some differences in the appropriate primary and secondary dendrites seemed to occur. Based on intracellular recordings it was shown (locust: Burrows, 1975; dragonfly: Simmons, 1980) that homologous MNs receive similar inputs from either a visual IN or the segmental wing stretch receptor. More information on the question of whether function determines the structure of identified neurones resulted from studies of leg MNs in Schistocerca (Wilson and Hoyle, 1978; Wilson, 1979a). The MNs innervating the appropriate muscles had been identified previously (Burrows and Hoyle, 1973). All three legs are used almost equally during walking, whereas the hindlegs are specialized for jumping. Homologous MNs are situated at a similar position in all three thoracic ganglia. Each cell has the same serially homologous neurones as neighbours. The only exception was seen for the pair of fast and slow extensor tibiae MNs. Apparently, for the metathoracic system the function of both neurones has been switched without any clear changes of morphological structures (position of soma, dendritic tree). Actually the central physiological connections between the elements is different when the pro- and mesothoracic systems are compared with that of the metathorax (Wilson, 1979b). From this it was inferred “that neuronal morphology is conservative even when a dramatic functional change has occurred” (Wilson and Hoyle, 1978).

20


Studies employing the large pterothoracic musculature resulted in a basic framework of MNs which are involved in the flight of locusts. Bentley (1970) used the technique of intracellular dye-iontophoresis (Procion yellow, Schistocerca) to identify these MNs; and lateral cobalt-backfills (with successive intensification; Timm, 1958; Tyrer and Bell, 1974) resulted in the demonstration of unexpectedly elaborated dendritic fields of these neurones (Tyrer and Altman, 1974). A comparison of the same neurone in different individuals (Chortoicetes terminifera) indicated that there is an overall similarity, though variation in the detailed pattern of branching occurred (e.g. number of primary branches, branching point along the neuropilar segment). However, since particular regions of the neuropile will be reached by the individual neurones, it is suggested that the found differences are probably of no, or only little, functional importance. Generally, the number and position of the somata connecting to homologous muscles of the mesoand metathorax is similar. For the MNs serving the dorso-longitudinal muscles (DLMs), the dendritic fields have also been described. The DLM is of special interest since its MNs are distributed between two ganglia (Neville, 1963). Four MNs are situated in the next anterior ganglion, ipsilaterally and a fifth MN is placed contralaterally within the ganglion associated with the muscle’s segment. This latter neurone innervates the most dorsal muscle bundle (Neville, 1963; Kutsch and Heckmann, unpublished; see Fig. 1A). The dendritic fields of both ensembles (meso- or metathorax) are similar, although the dendritic field appeared to be much less extensive for the metathorax (Tyrer and Altman, 1974). Since it is to be expected that homologous MNs serve homologous muscles one should, for a further generalization, prove a validity of this theory by checking all segmental muscles along the body axis. For the pterothoracic system (mesoand metathorax) the equivalence of muscles is rather obvious; however, the prothorax and the abdominal segments diverge in their muscle sets. Nevertheless, this does not rule out the possibility that for each muscle of these other segments there are homologues in the pterothorax (which does not mean that this assumption is also true for the reverse case). We have chosen to study the neural set which supplies the DLMs in locusts (Breidbach and Kutsch, 1990; Kutsch and Heckmann, unpublished). These muscles are present throughout all segments and seem to fulfil the criterion of “serial homologues” . Equivalences for other segmental muscles still need to be proven (Voss, 1905; Snodgrass, 1935; Xie et al., 1992). The whole DLM-complement is composed of three muscles of which one or both smaller ones degenerate in the pterothoracic segments during early imaginal life (Wiesend, 1957). The neural set is almost identical for all segments (see Table 1). There are several typical MNs as well as specific cell


21

FIG. 1 Photography of wholemounts (NiC12-backfills, intensified) showing the MNs supplying the DLMs of the pterygote Schistocerca (A) and the primary apterygote Lepisma (B). In both cases, the DLM of the metathorax has been chosen. The MNs split up between the mesothoracic (MS, via nerve 6 ) and metathoracic (MT, via nerve 1) ganglion. (A) arrowhead, indication of the larger contralateral MN; arrow, axon of the bilaterally exiting D U M neurone (soma out of focus), the median ipsilateral fibres of M T are of sensory origin. (B) straight arrow, soma of the large contralateral MN; curved arrow, soma of a large loop cell. (Modifed after Kutsch and Heckmann, unpublished (A), and Heckmann and Kutsch, unpublished (B).)

types, which are distributed between two ganglia for the muscle system of one segment. In Schistocerca, the number and position of neurones innervating the first abdominal DLMs is equivalent to the aforementioned results (Yang and Burrows, 1983). The only dissimilarity in the otherwise structurally conserved MN is seen in an EM analysis resulting in a much higher number of axons serving the relevant abdominal DLMs (Tyrer, 1971). The reason for this divergence is still unclear. Also for the neurones supplying the median internal ventral muscle (Yang and Burrows, 1982), an apparent homologous set can be identified for the thoracic segments (Steffens, unpublished).

TABLE 1 Identified MN types serving the DLMs in different segments of Schistocerca; adult and embryonal stage Muscle(s) DLMs Pro Adult (M 49 & 56) DLMs Meso Adult DLMs Meso Embryo 95% DLMs Meta Adult DLMs Meta Embryo 65% DLMs AS 7 Adult

Anterior neurone group Posterior neurone group Ganglion 1MN mMN sMN loop C1,3,6 pod0 Sum Ganglion contra C1,3,6 DUM cc Sum

SOG

3

2

3

1

?

1

9

PRO

2

?

1

1

4

PRO PRO MESO MESO A6

4 4

1 1

3 3

(1) 1

(1) 1

11 11

MESO MESO

2 2

(1) 1

1 1

1 1

5 5

4 4 3

1 1

3 5

(1) 1 1

(1) 1 (1) 1 1

(1) 2 1

11 14 7

META META A7

2 2

(1) 1 1

1 1 2

1 1 1

5 5 6

1

2

The MNs are characterized using several criteria. The neurones are split up into a group positioned in the next anterior ganglion and a group positioned in the (posterior) ganglion associated with the muscles. lMN, large motoneurones; mMN, medium sized motoneurones; sMN, small motoneurones; loop, neurone with a contralaterally looping neuropilar segment; contra, neurones with a contralateral soma position; podo, soma in a posterior-dorsal position; DUM, dorsal unpaired median neurone; cc, central cell; C1,3,6, apparent serial homologous neurones in both groups located contralaterally, their axons split up and exit via nerves N 1,3,6 (thorax) or N 1,2 and posterior connective (abdomen). Ganglion: SOC, suboesophageal; PRO, MESO, META, pro-, meso-, metathoracic; A6, A7, sixth and seventh abdominal. The typical number of cells for each type is given. ?, this neurone type has not been detected, though it might exist; (l), a single cell seen in only a few preparations. For the prothorax two muscles (nomenclature after Snodgrass, 1929) have been selected. (Modified after Kutsch and Heckmann (unpublished).)


23

These muscles are present from the first larval stage onwards (Wiesend, 1957; Bernays, 1972) and actually, the typical muscle pattern is reached near to 70% of embryogenesis (Xie et al., 1992). The whole neural complement (including the characteristic features of the different cells) is present in a 65% embryo (see Table 1); for one of the MNs the embryonic development has been followed demonstrating the differentiation towards typical adult structures reached at about 65% of embryogenesis (Steffens and Kutsch, 1992). A direct approach of comparing a few other muscles in both pterothoracic segments of locusts was carried out by Kutsch and Schneider (1987). This revealed a very close matching of the apparently homologous MNs in both ganglia. A more thorough description of almost all MNs in a grasshopper is provided by the studies of Siegler and coworkers (Siegler and Pousman, 1990a,b; Siegler et al., 1991b). Although these descriptions are restricted to the metathoracic segment, they represent a basis for a search for possible homologues in other segments. 3.1.2.1.2 Other MN-types. Typical MNs are characterized by their lateral, usually ipsilateral, and ventral soma position. They exhibit a dorsal dendritic field restricted to the appropriate hemiganglion. There are several other cell types which connect to the muscles (therefore they are also termed MNs). They might be involved in neurosecretion and usually show several deviating morphological characteristics. A few have been listed in Table 1. In the following we will restrict our comparative approach to the DUM neurones and the common inhibitory neurone. The DUM cells are characterized by a dorsal soma position; this soma is unpaired and gives off a single neurite which splits off to both sides, where a scarce dendritic field is elaborated. Such a group of up to nine neurones has been seen in all thoracic ganglia. They exit the ganglion where the axons split in up to four branches. In the 4th abdominal ganglion a single neuron was detected (Plotnikova, 1969). Other studies have mainly concentrated on number, distribution, and physiological aspects, usually restricted to the metathoracic ganglion of locusts (Crossman et al., 1971; Hoyle, 1978; Hoyle and Dagan, 1978; Heitler and Goodman, 1978; for recent studies concerning the effects of DUM cells on a flight muscle, see Whim and Evans, 1988, 1991). All DUM neurones are the progeny of the single median neuroblast in each segmental ganglion (Goodman and Spitzer, 1979; Goodman, 1982). For some time, an association of DUM neurons with the neuromodulator octopamine has been claimed, though for each neuron a direct proof has to be carried out (Evans, 1978, 1980, 1985; Agricola et al., 1988). Recent studies demonstrated that the group is not homogeneous. They are different with respect to cofactors of enzyme synthesis (Siegler et al., 1991a), the small local DUM neurones are stained by GABA-antibodies (Thompson and Siegler, 1991; Stevenson et al., 1992) and single abdominal DUM cells seem to contain FMRFamide-like transmitters (Ferber and Pfluger, 1992).

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Octopamine antiserum stains only a small population of the relevant cell groups (Konings et al. , 1988a), restricted to those projecting peripherally (Stevenson er al., 1992). Based on such immunocytological analysis it is quite obvious that there are similar neural candidates distributed along the different segments (see for instance table 1, in Stevenson et al., 1992); however, position and number of the somata vary, especially with respect to the thoracic ganglia, which, at the moment prohibit their unequivocal determination as “serial homologues”. Inhibitory MNs have been detected in the thorax of the locust (Pearson and Bergman, 1969; Burrows, 1973). There have been three cells described, all of which are common inhibitors. CI, serves 13 muscles while C12 and C13 have a more restricted peripheral field, comprising four muscles (see Fig. 14B). At least for all three thoracic ganglia, it is obvious that the equivalent cells (soma position, dendritic structures, etc.) exist (Hoyle, 1975; Watson et a f . , 1985; Hale and Burrows, 1985). This seems fully to conform with the assumption of serial homologues independent of segmental structure (e.g. pro- vs. metathoracic segment). Lang and Wolf (1992) describe the origin of the common inhibitors 1 and 3 as progeny of neuroblast 5-5 in the mesothorax of Schistocerca. Incidently, this specific neuroblast is missing for the abdominal segments 1-7 (Doe and Goodman, 1985). Therefore, it would not be expected to find these inhibitory MNs in these segments, though they might be present in the last abdominal ganglion. This is an interesting case for the potency of the analysis of early neurogenesis in the search for homologues. The progeny of the median neuroblast has been studied especially well, since, owing to its exposed position, it gives a fortunate opportunity to follow up the fate of identified neurones (cell lineage from serial homologues, number of divisions, etc.) (Goodman and Bate, 1981; Goodman, 1982). Each median neuroblast produces about 100 progenies in the thorax, while in the abdomen about 90 are produced of which, however, only half will survive. Under the argument that the cell lineage is identical for all segments, it would be expected, at least for the remaining 45 cells of the abdominal system, that there are serial homologues in the thoracic CNS. Actually, it is claimed (Goodman and Bate, 1981) that in the abdominal segments two MNs develop (DUMETi and FETi, stemming from different neuroblasts) which are “identified as homologous to their thoracic namesakes by lineage”. In the thorax, they serve leg muscles, while, in the abdomen, they grow towards the edge of the CNS and then die. Therefore, the initial programme of the segments is identical (+ production of a homologous set of cells) though the specific requirement of the segments (leg-bearing vs. no legs) renders these cells to be “obsolete”. 3.1.2.2 Phylogenetic considerations In the search for a possible common “Bauplan” of the Insecta segment one should compare the musculature and the appropriate neural set employing a rather wide range of species within


25

this animal group. This approach is supported by the knowledge that there appears to exist a common plan for neural development (Breidbach and Kutsch, 1990), from a more primitive insect, the grasshopper, to one of the most advanced, the fruitfly Drosophila (Thomas et al., 1984). Not only are the number of neuroblasts and their position similar for both insects but “the Drosophilu embryo is a miniature replica of the grasshopper embryo in terms of its identified neurones, their growth cones and their selective fasciculation choices”. Of all the possible muscle sets the dorso-longitudinal muscles are the most obvious ones to be compared throughout different species. Therefore, the appropriate structure of MNs has been studied in a wide range, from the “primitive” silverfish Lepismu to Drosophila (see Fig. 1B and Table 2). Taking into account the fact that the studies have been carried out in different laboratories, with slightly different techniques, it is not unreasonable to suppose that at least this neural ensemble is equivalent, suggesting they may represent a homologous set of neurones throughout the Insecta. As a further point of interest, it is apparent that structurally homologous motoneurones are not only identified in the adult stage, but that they are present throughout larval development. They even survive metamorphosis in Holometabola (Breidbach and Kutsch, 1990). This supports the idea that not only the neurones per se, but also their appropriate structures, are conserved irrespective of changes in the periphery and strategy of postembryonic development. Homologous MNs are also observed in a flightless grasshopper (Arbas, 1983b). In these creatures the DLMs degenerate in the pterothorax (Arbas, 1983a); leaving innervation and presynaptic structures along inappropriate targets (Arbas and Tolbert, 1986). With respect to the migratory locust the relevant homologous MNs occupy an equivalent position, though the soma sizes and probably also the dendritic fields, are reduced in the flightless grasshopper. One of the MNs supplying the extensor muscle of the tibia, the fast extensor-tibiae (FETi) MN of the metathorax, has been compared in different acridid and gryllid insects (Wilson et al., 1982). (A rather similar structure is apparent within each group (acridid vs. gryllid); the difference between both groups points to a specialization of the FETi in acridids towards a jumping function (see above, a similar argument for serial homologues) . For a fine control of head movements, the neck muscles are arranged in a seemingly sophisticated manner. Also, the innervation is complex with muscles sharing the same MNs. Honegger et al. (1984) have compared the relevant motor system of crickets with that of locusts. Any complexity of innervation per se is a disadvantage for analysis; however, as mentioned above, shared complexity involving identified units improves the possibility for a claim of homology. For almost each of the described MNs, a similar

26


TABLE 2 Comparison of type and number of MNs supplying the unisegmental DLMs of different insect orders with the innervation of a Chilopoda and Crustacea Systematic unit

split

iaMN

cpMN

DUM

loop

cc

C1,3,6

7

2

4 7 7

1 2 1 2 2 1

+ + + + + + ? +

(+)

+

(+I

+

(+>

+

+

? -

-

?

?

-

~~

Lepidoptera’ Dipterab Coleopterac Heteropterad Blattariae Saltatoria Odonatd Zygentoma Chilopoda Crustaceag

+ + + + + + + + + +

6 8 4

6 9

2

4

3

?

1 ?

+ 1 ?

? 1 ?

1 ?

? ? ?

+

4

The MNs have been classified using common features. split, neurones are distributed throughout two ganglia (next anterior and muscle segment associated); iaMN, ipsilateral somata, anterior neural set; cpMN, contralateral somata, posterior neural set; DUM, neurones with DUM-like morphology; loop, soma located contralaterally, close to the midline, neurite and neuropilar segment loops contralaterally before entering the ipsilateral nerve; cc, central cell; C1,3,6, neurone with soma in contralateral position, axon splits and exists via nerves N 1,3,6. +, at least one cell present; -, no cell present; ?, cell might be present, but has not been reported; (+), some indication for presence of this cell type. a Bornbyx, Tsujimura (1988, 1989); Drusuphila, Ikeda and Koenig (1988); Zophubas, Breidbach and Kutsch (1990); Dysdercus, Davis (1977); Periplaneta, Davis (1983); Hemianax and Aeshna, Simmons (1977); g Procambarus, Mittenthal and Wine (1978). Occasionally, some additional information from other authors has been included; information from our own work: Schistucerca (Saltatoria), Lepisma (Zygentoma), Lithobiur (Chilopoda). (Modified after Heckmann and Kutsch (unpublished).)

’

partner is found for both species. In cases where equivalent MNs are seen innervating differently arranged muscles it is argued that the muscles have changed their orientation and function; for a similar argument employing possible serial homologous muscles in crickets, see Bartos and Honegger (1992). If it turns out that the structures of the MNs are more “conservative” than their peripheral targets, then one could, in the future, use the MNs to search for homologous muscles (see also Paul, 1991). Among other aspects, this hypothesis requires embryological studies with early demonstration of the muscles (Myers and Ball, 1987; Xie et al., 1992) and their MNs (Whitington and Seifert, 1981; Myers and Ball, 1987; Myers et al., 1990; Sink and Whitington, 1991; Steffens and Kutsch, 1992). Another meaningful approach is to compare identified MNs within an insect group in which the periphery varies considerably. Phasmids are well suited for such an investigation. This insect order comprises a wide range with respect to the configuration of the wing system, from excellent fliers to


27

FIG. 2 (A) Flight position of members of the Phasmida. Sy, female of the winged Sipyloidea sipylus; Ca, female of the wingless Carausius morosus. (B) Camera lucida drawings of the two mesothoracic MNs supplying one of the major flight muscles (subalar muscle) in winged species. The dendritic fields of both species are rather similar, only in the central ganglionic region a reduction for Ca is seen when compared with Sy. For comparison the two subalar muscle MNs of a female Locusta (Lo) are also presented, exhibiting similar features, though the dendritic field is denser. Scale 100 ym. (After Kutsch and Kittmann (1991, and unpublished).)

28


reduction of the flight system and, ultimately, to unwinged species (for summary, see Kutsch and Kittmann, 1991). Correlated with the reduction of the wing system there is also a reduction of the major flight muscles. Even though, the appropriate MNs (number, position, dendritic organization) are very similar (Fig. 2). There are only minor differences pointing to some positive correlation between flight ability, soma size and extension of the dendritic projection (Kittmann and Kutsch, unpublished). Since homologous MNs are found for the locust, too, this aspect of central conservatism is substantiated compared with a more freely changing periphery.

3.1.3 Sensory neurones 3.1.3.1 Serial homology The process of reiteration of developmental processes for the generation of a segmented animal may also result in a repetition (homologous patterns) of peripheral nerves and sense organs. Recent techniques in genetics and staining have indicated such a repetition, especially well in embryos, from grasshopper to Drosophila (Jan and Jan, 1982; Campos-Ortega and Hartenstein, 1985; Bodmer and Jan, 1987; Hartenstein, 1988). For Drosophilu larva, several sensory organs have been described (summary in Hartenstein, 1988): sensilla trichoidea, sensilla campaniformia, sensilla basiconica, and chordotonal organs. At least for the thoracic and abdominal body, practically all these receptor types are present along each segment. Even though there are slight differences in position and number of individual receptors within one field when comparing the different segments (Campos-Ortega and Hartenstein, 1985), serial homology can be claimed up to the level of individual receptor cells (Hartenstein, 1987, 1988). The generation of the different receptor types occurs in a dorso-ventral and rostro-caudal temporal gradient, while the differentiation of the apparent homologous sensory neurones occurs more or less simultaneously along the embryo (Hartenstein, 1988). With respect to the larval sensory system it becomes apparent that all appropriate receptor organs are present in the late embryo. Towards the end of stage 15 (for definition, see Campos-Ortega and Hartenstein, 1985) all receptor types can be revealed (Hartenstein, 1988). Compared to the duration of embryogenesis, this period equals almost two-thirds of the total embryonic development. Receptors are generated by a complex differentiation of epidermal cells. It is still accepted that along the epidermis of each segment a gradient of some substances exists; such positional information may result in the generation of the specific receptor cells and their accessory cells, including the specific formation of the overlying cuticle (Gnatzy and Romer, 1984; Kutsch, 1989). Since the same cues appear to be repeated throughout the different segments similar (serial homologous) sensory structures will be generated repeatedly. One problem of early receptor genesis is to find a


29

path to the CNS along a variety of non-neural tissue. It turns out that the growth cones of the receptors use a given path initiated by specific pioneer neurones. Later, these neurones may be converted into sense cells (such as in Drosophila wings; Blair and Palka, 1985) or die during later stages of embryogenesis (pioneer of the hindleg receptor system: Kutsch and Bentley, 1987). The pioneer axons navigate towards the CNS along guide-post neurones (Bentley and Keshishian 1982a,b; Bentley and Caudy, 1983); however, other cues may also be utilized for which interspecific equivalences have been suggested, though it needs to be proven for each case (Palka, 1986; Lefcort and Bentley, 1987; Nardi and Vernon, 1990; Ruiz-Gbmez, 1990; Singer et al., 1992; Norbeck et a f . , 1992). For the locust, the pioneer cell (pair) has been detected at the tip of the antenna, in the mouthparts, for all legs and the cercus (Bate, 1978a,b; Shankland, 1981; Bentley and Keshishian, 1982a; Meier and Reichert , 1991). Therefore, this system, encountering serial homologous appendages, appears to be a convincing case of an identified homologous pair of neurones along the body axis. Unfortunately, a description of the process of pathfinding has mainly been restricted to the locust hindleg. Therefore, axonal navigation for the different appendages has not yet been determined; though, it is to be assumed that homologous processes occur along each route (see Meier and Reichert, 1991b). This conclusion is based on studies with antibodies against membrane proteins. Their analogous expression gives supporting evidence for the assumption of homologous navigation pathways in the appendages of different insect species. According to their studies, the pleuropods represent the basic pattern of an insect appendage (Meier and Reichert, 1991b). Similar pioneers have also been detected in other species (crickets, Edwards and Chen, 1974; cockroaches, Norbeck et al., 1992).

The ontogenesis of the leg receptor and internal sense organs has been described during grasshopper embryogenesis (Kutsch, 1989; Meier et af., 1991). It is apparent that within about two-thirds of embryogenesis the proprioceptors have reached their final appearance, and that the external receptor fields are present also, though the number of individual receptors will increase throughout larval life (Sviderskij, 1969; Altman et al., 1978). In the following section, knowledge on segmental homologies for adult animals will be reviewed. Although it is to be expected that the generation of the sensory system is reiterated along the segmental body, it is astonishing that there are not too many cases available (in which this specific aspect has been discussed). Hustert (1978) describes a pair of mechano-sensitive cells placed on each segmental sternite in a cricket (Acheta). This singularity gives an opportunity to visualize the central path of the appropriate primary afferent fibres. They extend ipsilaterally over several ganglia; the extension and branching fields in each ganglion are similar for all segmental inputs. Similar central branching can be shown for other segmental hairs (Hustert, 1985). Comparison of the complex central distribution (Hustert, 1978) also

30


indicates that the multipolar wing stretch receptor in locusts (Gettrup, 1962) may be homologous to the abdominal stretch receptors (pleural chordotonal organ; Hustert, 1974; Finlayson, 1976). In a classical series of investigations on two locust species, the similarity of the morphology and central arborization of internal and external receptors associated with the three thoracic segments and their legs has been shown (Braunig et al., 1981; Pfluger et al., 1981; Hustert et al., 1981). Within all three locust legs, at the femur-tibia joint, one chordotonal organ is described (Slifer, 1935; Usherwood et al., 1968; Burns, 1974) and a group of five multipolar sensory neurones (Coillot and Boistel, 1968; Williamson and Burns, 1978). Later, a strand receptor was added to the list of mechanoreceptors. It is a single cell neurone, with a soma positioned in the CNS (see later), connecting to a ligament associated with the femoral chordotonal organ. This sense cell is observed in all three legs (Braunig, 1985). There are slight differences in the route of the appropriate axon; though the peculiarity of this mechano-receptive system points to a serial homology of a further identified neurone. The discovery of proprioceptors in insects with their cell bodies positioned, unexpectedly, in the CNS (Braunig and Hustert, 1980) prompted a (positive) search of homologous cells for all three legs of locusts (Braunig, 1982a). Such a specific group of cells with central somata has been observed in a selection of orthopteran insects (Braunig, 1982b). A comparative study on two coxal sense organs, which can be associated with peripheral end organs within the coxa in a variety of pterygote insects (Braunig, 1985), should support the notion that homology of receptor systems and individual sense cells is the rule, not the exception. There are several receptor systems at the mouthparts which are equivalent with those of the thoracic appendages. Central projections show several features in common (Braunig et al., 1983). However, there are differences; as a caveat “intersegmental homology, if ever present, probably was strongly overruled by functional demand”. Owing to the general interest in acoustical communication there is a wealth of information concerning behavioural role, structure and function of auditory organs, especially for Orthoptera (for a synopsis, see Kalmring and Elsner, 1985). The position of the appropriate organ is mainly restricted to one segment: either in the pleural wall of the first abdominal segment (Caelifera) or in the tibia of both forelegs (Ensifera). This restriction to one segment might have prevented the search for possible serially homologous structures along the body. However, serial homology should be propounded. Friedrich (1927, 1928) investigated the subgenual organs in Tettigoniida, and he gave evidence for a homology of the organs in all legs. The organs of the middle- and hindlegs were almost identical, apparently being more representative of the original set in Tettigoniida. The development of tympani in the forelegs required a secondary reconstruction (e.g. increase in number of scolopidia, change in the structure of the membrana tectoria).


31

For a review on the peripheral nervous system, especially with respect to the foreleg containing the auditory organ of Tettigoniida, see Lakes and Schikorski (1990). In a detailed study (Lakes-Harlan and Mucke, 1989), the receptor fields and the nerve branches of a tettigoniid auditory organ have been described for several individuals within one species (Ephippigger ephippigger, Tettigoniida). The basic pattern is rather congruous; however, individual fibres could join the main nerve at different locations. This apparently, reflects the freedom of the periphery compared to the CNS and may give the opportunity for change in the periphery in comparison to the more conservative CNS. It becomes clear that during early embryogenesis an equivalent organ anlage is present in all three legs of Tettigoniida. Subsequently, prolonged cell invagination and migration mainly in the foreleg results in the observed structural differences. Also for the locust ear receptor array, serially homologous sensory organs could be shown by studying the developing embryo (Meier and Reichert, 1990; Fig. 3). In the thoracic segments chordotonal organs are present close to the wing hinge, whilst in abdominal segments pleural chordotonal organs are evident, the first one of which develops into the auditory organ (homology!). Cell migration, pathfinding and expression of a specific membrane molecule, seen in parallel for all segments, are additional hints for serially homologous structures and physiological processes. Furthermore, the tympana1 organ of noctuids can be homologized with the locust metathoracic chordotonal organ at the wing base (Yack and Fullard, 1990; see original statement, Wilson and Gettrup, 1963). A comprehensive treatise (Meier et al., 1991), also based on embryonic studies of a locust, demonstrated a serial homology of a specific nerve and its associated sense organs. Added to this is the discovery that a segmentally reiterated pattern also exists in Drosophila. Furthermore, for Schistocerca and Drosophila segmental sets have many features in common, suggesting a conservation of the peripheral nervous system of ancestral insects (Fig. 3). 3.1.3.2 Interspecific homology Some additional examples of interspecific homology will be considered in the present section for comparison with those already mentioned above. The advantage given by the single-celled wing stretch receptor neurone with its large axon has often prompted behavioural and physiological studies (e.g. Wilson and Gettrup, 1963; Burrows, 1975; Pearson et al., 1983; Pearson and Ramirez, 1990; Ramirez and Pearson, 1990). It has also been studied during locust ontogeny (Altman et al., 1978). The central paths and arborizations of both fore- and hindwing receptors are very complex, with little variation even at the level of fine detail (Altman and Tyrer, 1977b). A matching of both systems is apparent and the similarity in three locust species is obvious. Therefore, for this identified receptor cell not only a serial, but also an interspecific homology, is manifested.


32

Hemimeta bola Holometabola Schistocerca Thorax (T2 & T3):

Drosophila Thorax (T2 & T3):

dorsal: dcl. dc2. dc3. dh2

I:

Abdomen (A1-MI:

p

+-".

Abdomen (Al-A7):

darsalz

FIG. 3 Schematic representation of segmental sensory structures and their associated nerves for an embryonic hemimetabolan (Schistocerca) and holometabolan (Drosophila) insect. In each species serial homology is shown €or both pterothoracic (T2&T3) and for the abdominal segments (Al-A8); furthermore, interspecific homology is apparent. (Modified from Meier ef af. (1991).)


33

A dependence of the size of the foreleg crista acustica with the developmental status of a tympana1 membrane is also inferred by comparing tympanate and atympanate Tettigoniida (Schumacher and Houtermans, 1975; Lakes-Harlan et al., 1991). Peripheral and central projections in an atympanate form are more similar to the meso- and metathoracic segment favouring a more “primitive” system compared to the typical prothoracic arrangement of tympanate species. In a systematic comparison of Tettigoniidea and Acridoidea, it becomes obvious that for each group similar central projections and interactions with identified auditory interneurones exist (Kalmring et al., 1985). However, within the so-called “homologous” neuropile of locusts and bushcrickets drastic differences in the wiring diagram occur (Romer et al., 1988). These results, demonstrating differences in structure and connections within a system of appropriate functional significance, represent a typical case of “analogues”. Homologues should not be expected, since both auditory organs are not considered to be homologous structures. The apparent similarity of the peripheral nervous branching pattern and the arrangement of the sense organs of the wings have been used to gain an understanding of the close relationship among species and among certain orders, as well as to compile a possible phylogenesis of insects, from Orthoptera to Diptera (Fudalewicz-Niemczky, 1958, 1963, 1968). 3.1.4 Interneurones (INS) The majority of neurones in the nervous systems of insects are interneurones. At present, there is no generally accepted scheme for classifying these interneurones (Weevers, 1985). Using gross morphological terms we can distinguish segmental and plurisegmental interneurones, according to whether their arborizations are restricted to one or to several ganglia (Pearson, 1978). Immunohistochemistry has contributed further criteria for a classification of neurones. 3.1.4.1 Ventral nerve cord (VNC) Most evidence for a comparative analysis is found in the ganglia of the VNC. We will first describe evidence for homologies of neurones gained by conventional neuroanatomy and comparative physiology (Burrows, 1973, 1985; Siegler, 1984; Rowel1 and Reichert, 1991). Thereafter, we will describe homologies of immunohistochemically characterized VNC interneurones. However, evidence for homologous brain interneurones is scarce, both in respect of serial and phylogenetic homologies. It is supposed that neuropile regions, such as the central complex, the mushroom bodies or the antenna1 lobes, are homologous structures in insects (Bretschneider, 1914; Hanstrom, 1928a; Bullock and Horridge, 1965; Strausfeld, 1976; Mobbs, 1985). According to the interpretation of mandibulate brain morphologies, as

34


provided by Strausfeld et al. (1984), it is doubtful whether one has to integrate these neuropile areas into a common scheme of the mandibulate brain. The orthopteran nervous system provides one of the established invertebrate model systems in which to study VNC processes in the control of motor behaviour and integration of sensory information (Huber, 1983). The basic neurohistology of this system is well established, providing a detailed reference for a description of the positions of identified neurones in different VNC ganglia (Tyrer and Gregory, 1982; Pfluger et al., 1988). With regard to motor systems, most interest has been directed to the flight system of the locust. Different studies have tried to map the interneurones involved in flight control (Robertson et al., 1982; Robertson and Pearson, 1983, 1985; Burrows, 1985; Rowel1 and Reichert, 1991). Relevant interneurones were found to be distributed within the three thoracic and the first three abdominal ganglia (Robertson and Pearson, 1983). These neurones could be divided into three organizational categories: members of two serially homologous groups controlling either the fore- or hindwings, unique individuals with no known homologues in other ganglia and members of a set of serial homologues in the metathoracic and first three abdominal ganglia. Robertson and Pearson (1983) described serial homologues by morphological and physiological criteria: location of the cell soma, path of the neurite, neuropile segment, major dendritic branching pattern and indication of similar responses to wind stimulation and similar activity patterns during flight sequences. One interganglionic interneurone, the interneurone 201, is present as a serial homologue in the pro- and mesothoracic segment. It connects the posterior dorsal region of the proand mesothoracic neuropile with the most dorsal neuropile in the anterior two-thirds of the meso- and metathoracic ganglion, respectively. This neurone is found also in both the first and the second abdominal ganglion, where it possesses contralateral projections. Four sets of neurones were described as serial homologues in the metathoracic and the first abdominal ganglia (interneurones 501, 504, 401) or in the first abdominal ganglion (interneurone 503). Morphologically, the interneurones 501, 503, and 504 share a common relative position of their somata in the dorsal posterior quarter of the ganglion, a contralateral projection of their neurite which extends anteriorly, and four main projection regions in the dorso-lateral, the ipsilateral median, the contralateral dorsal median and the lateral neuropile. The neurite of interneurone 401, whose soma is ventrally medially located, ascends ipsilaterally forming ipsilateral projections in a region overlapping with those of the lateral projections of interneurones 501, 503 and 504. Robertson and Pearson (1983) discuss the evolutionary consequences of equivalent physiological patterns of identified serially homologous neurones. A question raised is what is the selective pressure due to the evolution of the flight motor system? In their opinion fusion of the first three abdominal


35

ganglia with the metathoracic ganglion occurred to reduce the conduction time between the components of the flight motor system. Remarkably, in spite of gross changes in ganglionic morphology, the principal shape of serially homologous neurones remains unaffected. Even such a functional “need”, if it ever existed, appears to be insufficient to transform the structural characteristics of the respective cells. Such persistence of structural characteristics of identified neurones irrespective of gross changes in the functionality is even more evident when comparing flightless and flying locusts (Arbas, 1983a). In his study, Arbas (1983~)found a close correspondence in the morphology of two interneurones, the DCMD, the descending contralateral movement detector, and the TCG, the tritocerebral commissure giant interneurone. They play a role in flight of locusts, but have counterparts in the flightless Mexican grasshopper, Barytettix psolus. By morphological characteristics these were identified as homologues. Apparently, with respect to the neurones in Schistocerca gregaria, the loss of flight ability in Barytettix has almost no effect on the structure of the TCG. There is a slight correlation with changes in the overall variability of the shape of the dorsal branch of the DCMD, which also shows a reduced area of innervation. However, these changes are close to the degree of variation found in an analysis of the variabilty of the morphology of the locust DCMD interneurone itself (Pearson and Goodman, 1979). Altogether, these examples demonstrate the structural persistence of homologous neurones irrespective of functional changes in closely related taxa. For Orthoptera, INS reflecting the “Gestalt” of the locust TCG have been found in the cricket, Gryllus campestris, and the mantid, Spodromantis linealoa (Bacon, 1980). The criteria by which these INS were identifed to represent homologues were: position of the cell body, projection pattern and size of the axon, dendritic arborization in the deuto- and tritocerebrum and projection sites in the suboesophageal and thoracic ganglia. However, the physiological characteristics of these cells were different. In the locust the TCG is very sensitive to stimulation of wind-hairs. It responds to antennal movements and shows slight activity correlated with light-on and light-off (Bacon and Tyrer, 1978; Bacon and Mohl, 1983; Mohl and Bacon, 1983). In the cricket the TCG homologue seems not to convey head hair input; it shows strong responses to head movements and produces a few spikes at light-on and light-off. In the mantid, there is also no head-hair input, the TCG homologue shows only small responses to head movements and a few spikes at light-on. Interestingly, while the magnitude of responses to antennal movements differs greatly in these three species, the direction of optimum stimulation (downward) is identical. Bacon (1980) discusses this as a criterion for homology. There are differences in the neural architectures of these TCG homologues; however, their principal structures are remarkably conserved. There are differences in the physiological reactions of these

36


neurons, which are irrespective of their structural invariance within the Orthoptera. Structural equivalence was also reported for auditory interneurones in the metathoracic ganglion of Chorthippus biguttufus and locusts (Locusta migratoria and Schistocerca greguria) (Romer and Marquart, 1984; Stumpner and Ronacher, 1991). Structural sameness was found in a set of 23 neurones. Three neurones are nearly identical in the position of their somata, the projections of their axons and in the distribution and structure of their side projections. These neurones exhibit similar physiological characteristics during processing of auditory signals. Apparently, information processing in the thoracic auditory neuropile is mediated by homologous neurones in locusts and grasshoppers (Ronacher, 1990). Thereby, structural characteristics of the respective interneurones are evolutionary conserved up to the degree of their fine arborization patterns (Stumpner and Ronacher, 1991). Some of these apparent homologous interneurones are restricted to the thoracic ganglia, while several others (the ANs) ascend to the sub- and supraoesophageal ganglia. One such ascending neurone, the G-IN is known to terminate in the lateral protocerebrum (Rehbein, 1976). Such neurones are reported to comprise a part of the auditory system for locusts (Rehbein, 1976), and for the moth and the fruitfly (Thomas et a f . , 1984). The neurogenesis of these cells is comparable in locusts and Drosophila (Thomas et al., 1984). In the latter species serial homologues are detected which are not involved in the processing of auditory information. Equivalent, homologous (G-like) INS are also present in the meal beetle, Tenebrio molitor, in the metathoracic and in the first four abdominal ganglia. In this beetle, which lacks a tympanum, these neurones are not involved in auditory information processing. They are already present in the VNC of the larva and persist after metamorphosis without major alteration of their structure (Breidbach, 1987~). Embryogenesis of the locust mesothoracic G-neurone and its segmental homologues (metathoracic and first abdominal segment) have been studied in more detail. Pearson et ul. (1985) established that their lineages originate from the neuroblast 7-4, the most lateral of the posterior row of neuroblasts (Fig. 4A,B). The principal morphology of the mesothoracic G-neurone and its serial homologues is already present in the 60% embryo and is maintained throughout further development (Fig. 4C). Therefore, the principal structure is established before these neurones become functionally active. It was shown that these serial homologues demonstrate significant differences in the organization of their inputs and outputs (Pearson et a f . , 1985). The authors argue that without an analysis of the embryonic lineage, homologies would not have been detected, solely based on morphological similarities. Segmental homologues apparently can be radically transformed structurally; they may have different physiological properties, and they may even die in different segments. Thus, according to this study, morphological


37

similarities may be sufficient to determine serial homologues, but are not sufficient in an identification of all possible homologues, for which a reference to embryology is required. In an analysis of the synaptic connectivity of the progeny of the 7-4 neuroblast in the meso- and metathorax and in the first abdominal segment of the locust, it has been outlined that all these cells receive synaptic input from a common presynaptic interneurone. Thus, the resulting neural network comprises postsynaptic cells with the same developmental lineage (Boyan, 1992). Detailed analyses of interactive processes in the developing insect nervous system indicate that serially homologous neurones develop due to a similar pattern in the VNC (Thomas et al., 1984; Bastiani et al., 1985). Thus, even cell interactions are formed due to a serially homologous mode. This means that serially homologous neurones contact serially homologous postsynaptic cells. Such contacts are a prerequisite for a proper outgrowth of the respective neurones. It is mediated by growth cone guidance via cell-cell interaction (Shankland et al., 1982). Accordingly, the reiterated shape of a serially homologous neurone is the result of a repetitive and equivalent programme of cell-cell interactions (Thomas et al., 1984). Such interactions remain effective even in later postembryonic development, where they allow the establishment of the structure of newly developing neurones (Sanes et al., 1976). Even structurally rather similar neurones, such as serotonin immunoreactive intraganglionic neurones (see below), maintain their complex similar dendritic arborization patterns in the metamorphosing meal beetle as a result of stereotyped cell-cell interactions (Breidbach, 1987d). Therefore, it is not surprising that structural variation of serial homologues is limited in most cases (Wilson and Hoyle, 1978; Burrows and Siegler, 1984; Burrows and Watkins, 1986), and it is independent of different functional surroundings. As soon as the population of neurons surrounding a serial homologous neuron is changed, owing to different developmental programmes in certain neuromers (depending on death of neural progenies, which prevents possible interactions with others), then the structure of the persisting serially homologous neurones may show striking differences. For example, the homologues with the locust H-cell are the progeny of the same precursor cell in different segments; they can either survive or die, and, in the former case, can develop different morphological and physiological properties (Bate et al., 1981). In consequence, it is suggested that similar serially homologous neurones in the adult stage are the result of serial homologous developmental steps, based on repetitive cell-cell interactions. If such programmes are stereotyped then it is not surprising, that serially homologous neurones are rather similar in different segments, even though they may perform different functions due to additional processes (Breidbach and Kutsch, 1990). Some examples are given for the DUM-cells and for ascending and descending neurones in various species (Braunig, 1988; Pfliiger and Watson, 1988; Ronacher, 1990; Breidbach, 1991).

38



39

The escape behaviour of cockroaches is mediated by a network consisting of wind-sensitive cercal receptors and giant interneurones which ascend to the thoracic ganglion where they excite the relevant motor units. Initially described for the cockroach (Roeder, 1948; Daley et al., 1981), this system has also been identified in other Orthoptera (Boyan and Ball, 1986; Boyan et al., 1989). The system appears to be a primitive one, since it also exists in the Thysanura (Edwards and Reddy, 1986). The system consists of sensory neurones of cercal filiform hairs. These project into the terminal fused ganglion ending in contact with a cercal glomerulus, formed by the densely interwoven arborizations of giant- and non-giant interneurones (Boyan et al., 1989). Because of the size of their ascending axons, the giants attracted early interest (Roeder, 1948). In the cockroach, in which the terminal ganglion is a fusion of five abdominal neuromers, seven bilateral pairs of such giant interneurones are present, with axon diameters between 15 and 30 pm (Daley et al., 1981). Comparative analysis of this system includes locusts, crickets and praying mantis (Fig. 5 ) . In the cricket, Jacobs and Murphey (1987) have summarized the segmental origins of the giant neurones during ontogeny. They have established that all of the giants derive from three segmentally repeated clusters of cells. It appears that the giant interneurones in other orthopteroid insects are also formed owing to this pattern. The somata of the giants are found in the ventro-median cell body group of the 9th, 10th and 11th neuromeres and in the lateral cell body group of the 9th neuromere (Boyan et al., 1989). Based on structural criteria the giant neurones found in these ventro-medial groups appear to be homologous to the respective neurones (&2a, 9-2a and 10-2a series) in crickets (Acheta; Jacobs and Murphey, 1987). There are enormous differences in the extent of development of the giant fibre system among extant Orthoptera, in absolute terms, and in relation to the size of other interneurones (Boyan and Ball, 1986). Thereby, the giant interneurones of the mantid seem to be more similar to those of the cockroach, than to those of either cricket and locust (Mendenhall and Murphey, 1974; Daley et al., 1981; Hue, 1983; Boyan and Ball, 1986; Boyan et al., 1989). Such structural

FIG. 4 Lineage of the mesothoracic G-neurone and its segmental homologues ( B l , B2) in the metathoracic and the first abdominal ganglion. (A) Pattern of neuronal precursors (61 neuroblasts; 30 NBs per hemiganglion, arranged in seven rows, and a single median NB (MNB) for a single segment. (B) The G-neuron arises from NB 7-4 (drawn in black). The G-cell homologues develop from a twin of a ganglion mother cell arising from neuroblast 7-4 in the appropriate ganglion. (C) Camera lucida drawings of the G, B1 and B2 neurone in adult Locusta migratoria; the Gneurone is located in the mesothoracic ganglion, the B1-neurone is located in the metathoracic neuromere, and the BZneurone is located mainly in the first abdominal neuromer, which is fused with the metathoracic neuromere in adult locusts. (Modified from Pearson et al. (1985).)

40

W. KUTSCH AND 0. EREIDEACH

similarities are difficult to interpret with regard to the gross morphology of the terminal ganglia. Differences for the various Orthoptera are due to different degrees of fusion of the last abdominal ganglia. In such fused neuropile, it might be difficult to decide whether similarity of neurones can be interpreted by constraints imposed upon the elements by surrounding cells or by true cellular homology. Owing to a comparative analysis (Boyan and Ball, 1986) based on a study of axonal pathways and the arborization patterns for individual cells, a homology has been assumed to exist. The equivalences are: the mantid lateral cell types 2 and 3 with the cricket 8-1 (MGI) and 9-1 (LGI) (Murphey et al., 1976, 1977) and the cockroach GI2 and GI3 (Daley et al., 1981; Hue, 1983); the cricket 10-2 (Mendenhall and Murphey, 1974) with cockroach GI6 (Daley et al., 1981) and the mantid cell type 7 (Boyan and Ball, 1986), the cricket cell 1&2 (Levine and Murphey, 1980; which is different from 10-2 in Mendenhall and Murphey, 1974) with cockroach GI5 (Daley et al., 1981) and the mantid cell type 5 neurone (Boyan and Ball, 1986). Beyond these homologies at the single cell level, the whole giant fibre system of orthopteroids reveals common characteristics. These comprise major pathways in common (ascending tracts and position within the commissures), the number of corresponding axons and their relative positions (Boyan et al., 1989). After all, this system seems to be highly conserved in evolution and is likely to be constituted-at least in parts-of identified homologous neurones. 3.1.4.2 Suboesophageal ganglion (SUG) Structural similarities detected for a whole neuropile area of the VNC can also be stated for the suboesophageal ganglia of different pterygote species. As described above, motoneurones serving the prothoracic DLMs in locusts (Schistocerca gregaria and Locusta migratoria), and beetles (Zophobas morio and Tenebrio molitor) correspond both in number and structural characteristics. This is also true for those motoneurons that are found in the suboesophageal ganglion (Honegger et al., 1984; Breidbach and Kutsch, 1990). Such correspondence is also found for serotonin immunoreactive, FMRFamidelike immunoreactive (Fig. 6), proctolin immunoreactive and crustacean cardioactive peptide (CCAP) immunoreactive interneurones in the suboesophageal ganglion of different insects (see below). A comparison of the principal features of suboesophageal descending neurones in Orthoptera (Kien and Altman, 1984; Kien et af., 1990) indicates some basic similarities with those described for the beetle Tenebrio molitor (Breidbach, 1991). The dendritic projections in the suboesophageal neuropile occupy corresponding areas in the beetle and locust (Kien et al., 1990). Likewise, several of the neurones described in detail for Locusta have their structural counterparts with Tenebrio neurones: SD2 in Locusta corresponds with a single neurone in the cluster vc2 of Tenebrio (Breidbach, 1991); SD 12 (Locusta) corresponds with neurones of c12 and m2 in Tenebrio; SD 22 (Locusta)


41

FIG. 5 Ascending neurones in the terminal ganglia of different Orthoptera; camera lucida drawings of the ganglia, wholemounts, seen from dorsal. (A) Locustu migruforiu, fibres leave the ganglion via the dorsal median (DMT) and the lateral dorsal tract (LDT), the abdominal neuromers 8-1 1 are indicated (8-1 1). Asterix points to a non-giant cell of the ventral median cell body group (VMCBG) of the 8th neuromer, which is a putative homologue of the giant interneurones 1, 2, 3 and 4, respectively (1-4). Scale bar 50 wm. (Modified from Boyan et al. (1989).) (B) Praying mantis, Archimantis sp., serial arrangement of identified larger cells (2-11) within the terminal ganglion. Scale bar 100 pm. (Modified from Boyan and Ball (1986).)

corresponds with A3 in Tenebrio; the group of SD 27-31 (Locusta) corresponds with a cluster of ipsilaterally descending neurones in Tenebrio. Correspondence is evident with regard to the principal structural features of the appropriate neurones, for example soma, location, sites of descending projection, and topology of dendritic projection (Kien et al., 1990). Mappings have not yet revealed the complete set of descending suboesophageal neurones in any of the species. Therefore, any possible homology can only be inferred on the close similarities of a few structural features. In any case, we should be aware that such similarities occur in phylogenetically rather separated species, namely locusts and beetles (Hennig, 1969).

42


FIG. 6 Serial homologous serotonin immunoreactive INS in the suboesophageal ganglion of the meal beetle, Tenebrio molitor, larva. (a) Camera lucida drawing, indicating a reiterated pattern of S1, and L immunoreactive neurones in the three suboesophageal neuromers (I, 11, 111); note change of position of the S2 soma in the different neuromers. (b,c,d) Horizontal sections through the suboesophageal ganglion demonstrating variation of the positions of serotonin immunoreactive neurones; levels of sections are indicated. Scale bar 50 pm. (From Breidbach (1991).)

For several of the suboesophageal descending neurones serial homology is assumed (Breidbach, 1991). It has not yet been possible to extend such serial homologues into the supraoesophageal ganglion, for which segmental organization is far less obvious, compared with the VNC ganglia. 3.1 -4.3 Supraoesophageal ganglion (SOG, brain) In the insect brain, somata, primary pathways and dendrites of descending neurones are arranged as discrete clusters, which cannot yet be associated with any of the


43

brain neuromeres. Even an analysis of the distribution of embryonic (Campos-Ortega and Hartenstein, 1985) and postembyronic neuroblasts (It0 and Hotta, 1992) did not allow the identification of segmental units in the developing brain, in contrast to the results obtained with the VNC (Bate, 1976; Truman and Bate, 1988; Truman, 1990; Prokop and Technau, 1991). In the developing embryonic and postembryonic brain of Drosophilu four mushroom body neuroblasts are prominent, placed medially and laterally from the calyces of the future mushroom bodies (It0 and Hotta, 1992). In the meal beetle, Zachaus (1992) identified four neuroblasts associated with the mushroom body. They have a similar location and show similar activity periods, compared with those identified in Drosophila. For Tenebrio, it was shown that the postembryonic neuroblasts are identical to the embryonic neuroblast associated with the mushroom body. For Tenebrio, they were found to be active from the early embryo (40% of embryogenesis) to the end of adult life (Zachaus and Breidbach, 1991; Zachaus, 1992). It is suggestive, that these four mushroom-body associated neuroblasts are true homologues for the two holometabolan insects, Drosophilu and Tenebrio. Consequently, this implies a common mode of development for at least a major part of the holometabolan mushroom bodies (It0 and Hotta, 1992). Thus, apart from structural criteria we have an additional (embryological) cue to homologize this part of the insect brain. This observation is of importance in view of a comparison of this brain region in arthropods, in general. In addition to enumerating certain structural criteria, we would define the tnushroom body neuropile as a neuropile, that should be, at least in part, derived from the proliferation of the homologues described for Drosophila and Tenebrio. Using the above information we gain criteria for only one subregion of the insect brain neuropile. However, it has not yet been possible to trace the supraoesophageal neuromere to which these neuroblasts belong. Traditionally, the insect brain is divided into three segments (Bullock and Horridge, 1965): 1. The tritocerebrum, which receives sensory input from the labrum and the stomatogastric system (incidently Chaudonneret (1987) provides some evidence, that this brain part is composed of two neuromers). 2. The deutocerebrum, which receives sensory input from the antennae. 3. The protocerebrum. Based on the expression pattern of segmental polarity genes (engruiled and wingless) during embryonic development of the larval head, SchmidtOtt and Technau (1992) give evidence that the head of Drosophilu consists of the remnants of four pregnathal and three postgnathal segments. All contribute cells to the neuromeres of the CNS. Accordingly, these data support the theory that there are four supraoesophageal neuromeres. The four patches of engrailed expression contribute to four pregnathal neuromers as follows: one spot is attributed to the intercalary neuromere, which may

44


represent both of the two neuromeres claimed by Chaudonneret (1987); one spot is attributed to the antennal segment, the former deutocerebrum (Bullock and Horridge, 1965); two further spots are associated with the protocerebrum, one of these is attributed to the ocular segment, to which the optic lobes fuse secondarily (Campos-Ortega and Hartenstein, 1985) and the other is attributed to the labral segment. These four segments are aligned in an S-shaped arrangement. These data fit with other comparative data. Frequently, for Insecta and Crustacea two pairs of coelomic cavities have been found anteriorly to the antennal segments (Wiesmann, 1926; Miller, 1940; Nair, 1949; Sharov, 1966; Rohrschneider, 1968). In his comparative analysis of embryogenesis, Wada (1966a,b,c) found seven segments for the head of Saltatoria (Tachycines). In accordance with these data, Schmidt-Ott and Technau (1992), suggest four basic brain segments: the ocular, the labral, the antennal and the intercalary segment. The authors suppose that these segments constitute the basic plan of the Mandibulata brain. Extending our knowledge about the basic scheme of neuroembryogenesis of the VNC of mandibulates (Pate1 et al., 1989a; Whitington et al., 1991) suggests that the head segments also seem to develop according to a general scheme. This may allow a comparative embryonic analysis, which is essential for an accurate definition of true homologies among different mandibulate groups, maybe even including the Chelicerata (Holmgren, 1916; Hanstrom, 1928a). It is difficult to relate insect brain architecture with the general scheme of an insect ganglion from the VNC (Weevers, 1985). This is mainly due to the fact that in the supraoesophageal ganglion serial homologies are difficult to demonstrate. As in the thoracic ganglia, within the brain interneurones also span over large neuropile areas and may arborize throughout the depth of the neuropile. In the thoracic segments it is possible to associate the complicated morphology of the neurones with the general scheme of a segmental ganglion. So far, a similar approach appears to have failed for the insect brain. Breidbach (1990~)has tried to uncover serial homologues in the insect brain, using ascending projections of serotonin immunoreactive neurones as markers (see below). One typical feature of the these neurones is their serial homology within the VNC (Taghert and Goodman, 1984; Tyrer et al. 1984; Hustert and Topel, 1986; Rheder et al., 1987; Breidbach, 1987a,b; Cantera and Nassel, 1987; Haeften and Schooneveld, 1992). There are two types of serial homologous immunopositive neurones (four bilateral symmetrical neurones), an ascending and local type of neurone (Fig. 7). Both cell types exhibit contralateral projections. Their ascending projections form a fascicle. It is possible to follow the fascicles of ascending axons into the brain and to depict those serotonin immunoreactive neurones which match to the ascending type of thoracic serotonin immunoreactive neurones. Breidbach


45

(1990~)proposed serial homologues of eight serotonin immunoreactive brain neurones with the four neurones of the VNC ganglion. Four of these neurones form part of the tritocerebrum, whereas only two are part of the antenna1 neuromere. Two neurones (MPN), observed in the protocerebrum, are other candidates for possible homologues, putatively forming part of the ocular brain segment. This set of serotonin brain interneurones proved to be conserved in insect phylogenesis (Breidbach, 1990~).Therefore, they may establish a reliable marker system to correlate different neuropile substructures of different insect brains, and may be used for an identification of brain neuropile homology in different species. Owing to the enormous diversity of cell types in the insect brain, it becomes very difficult to homologize certain neurones in different species based solely on morphological criteria. Incidently, only for the housefly Musca domestica is a detailed map of the brain available (Strausfeld, 1976). There are some histological accounts concerning Apis mellifera (Mobbs, 1985) and Drosophila (Power, 1937; Hanesch et al., 1989). Some scattered information exists about several neural elements of the beetle’s brain (Wegerhoff and Breidbach, 1992). However, these fail to give an appropriate reference allowing homologization depending on morphological criteria. Studies may become more prosperous due to the introduction of

FIG. 7 Serial homology of VNC, deuto- and tritocerebral serotonin immunoreactive neurones. Scheme of projection pattern of one half of the total set of neurones in the thoracic (TG), suboesophageal (SU) neuromeres and of putative serial homologues in the brain neuromeres (BRAIN) of Tenebrio molitor. AN, ascending neurones; d, deutocerebrum; DN, deutocerebral neurone; LN, local IN; MPN, median prothoracic neurone; p, protocerebrum; t, tritocerebrum; TNI,*tritocerebral neurones 1,2. (From Breidbach (1990b).)

46


immunohistochemical methods. Up until now, information is available only for a few insect groups. For Orthoptera, which comprises the standard system for the insect neurobiologist, no detailed information is available concerning the neuropile brain architecture (Williams, 1972, 1975). Formerly, comparative anatomy was centred on a description of the prominent neuropile areas in the insect midbrain (for some remarks on the visual system which includes the optic lobes, see below). Two midbrain neuropile areas received special interest, the mushroom bodies and the central complex (for review, see Gupta, 1987b). They were considered to be the most important brain areas in the regulation of complex behavioural patterns in Mandibulata (Erber et al., 1987). For the insect brain, homology of these neuropile regions has become established. However, a problem is whether these neuropile regions are true homologous structures for the Mandibulata or even for the Arthropoda, in general (Breidbach, 1992a,b). A more general view on an arthropod brain organization requires a reliable scheme of insect brain neuroarchitecture, which-unfortunately-has not yet been established (Mobbs, 1985). Based on morphological criteria, Strausfeld et al. (1984) give an account of the main brain neuropile centres. They suggest the following division: 1. The protocerebrum comprises the most anterior preoral neuropile. It will include the mushroom bodies, their laterally and medially surrounding neuropiles and the central body complex. 2. The deutoceberum is mainly composed of the antennal lobes, including the antenno-glomerular tract, the optic lobes and their associated tracts and neuropiles. Criteria for this assumption are found by an analysis of the structure of sensory projections from the sensory neuropiles in the brain. Thus, such a view could easily be combined with the embryonic analysis discussed above. There, it was distinguished between an ocellar and a labral segment, dividing the deutocerebrum (sensu Strausfeld) into two subunits, the antennal and the ocellar segment. Wegerhoff and Breidbach (1992) found that all the major midbrain optic pathways of the adult are present in the larva. They exist before the optic lobes are developed. Whether such a view may result in a better understanding of the tract organization in the insect brain needs more detailed morphological and developmental studies. For the Diptera interspecific homology has been established for one cell population of the midbrain: the giant fibre pathway. This is composed of intersegmental descending neurones that link the insect brain to the thoracic ganglia. These neurones exhibit dendrites in a specific region of the lateral deutocerebrum (sensu Strausfeld). The appropriate dendrites reach a multiglomerular complex formed by an ensemble of sensory neurones (Milde and Strausfeld, 1990). For the blowfly, these authors describe a cluster of eight descending neurones, characterized by a giant descending neurone. They receive primary mechanosensory afferents from the antennae


47

and the terminals of wide- and small-field retinotopic neurones originating in the lobula. The giant fibre terminates in the thoracic ganglia. This giant descending neurone is also identifiable in Drosophila (Power, 1948; Tanouye and Wyman, 1980; King and Wyman, 1980; Koto et al., 1981) with apparently similar functions (Nachtigall and Wilson, 1967; Milde and Strausfeld, 1990). It is found in Musca domestica (Coggeshall et al., 1973; Strausfeld, 1976; Strausfeld and Nassel, 1980) and in Sarcophaga bullata (King and Valentino, 1983). Based on structural features (soma position, dendrite structure, relative diameters of the axons, pathways, neurones contacted) King and Valentino (1983) claimed these neurones to be homologous. There are no structural variations apparent including the set of associated thoracic neurones. When compared with Drosophila, Sarcophaga shows a slight variation in the course of the giant descending neurone in the anterior thoracic region and a more pronounced branching pattern of the thoracic endings of this giant neurone. The analysis of the associated neurones in the thoracic ganglion also included the peripherally synapsing interneurone and the DLM MNs. For the whole set, neural homology for the three flies was claimed with only small differences, which can be attributed to the varying size of the three species (King and Valentino, 1983). This system seems to be comprised of homologous elements in all muscoid Diptera (Schizophora). Based on this assumption it was concluded that this pathway has been lost secondarily in one muscoid, Glossina morsitans (King, 1983). Structurally, the general scheme of such a descending pathway appears to be quite similar, even with regard to more distinct insect species. Structural features, as outlined for the dipteran descending brain interneurones (Strausfeld et al., 1984) can also be detected for the beetle Tenebrio molitor (Breidbach, 1989). There are similarities of descending neurones of group I1 in Tenebrio with PI 215 of the locust (Kien and Altman, 1984). The grasshopper TCG interneurone (Bacon and Tyrer, 1978; Bacon and Mohl, 1983), exhibits several conformities with neurones of group IV in Tenebrio. Presently, only structural similarities have been demonstrated. Nevertheless, they indicate a structural identity in the formation of descending neurones of three separated insect groups, Orthoptera, Diptera and Coleoptera. 3.1.4.4 Zmmunoreactivity The introduction of immunohistochemical methods may establish additional criteria for the homologization of neurones. Rowel1 (1976) reviews studies in which such techniques were applied to identify neurosecretory cells in the insect nervous system. Panov (1982) describes putative homologues for the ventral neurosecretory cells of Orthoptera. Elofsson (1972) and Aramant and Elofsson (1976a,b) used the Falck-Hillarp technique for a description of putative homologous brain structures in Insecta and Crustacea. However, no study allowed the identification of single neurones. Therefore, a reliable and detailed

48


descripton of the fine structure of the nervous system remained open. However, descriptions of cell populations that are immunoreactive to certain neuroamides or neuropeptides has rendered a detailed analysis of the morphologies of specific neurones possible and surmounted former attempts based on histochemical and conventional anatomical techniques. In addition to structural and histological criteria, immunohistochemistry gives additional criteria to identify homologous neurons. Supported by a thorough neuroanatomical description these data can provide reliable evidence of neural morphology. Furthermore, it is possible to study the ontogeny of such neurones and describe their lineage during embryogenesis. Thus, we can establish progenies and determine their neuroblasts (Taghert and Goodman, 1984). In this respect, the segmental structure of the nervous system is highly advantageous. Immunohistochemical studies allow identification of serial homologues. Since an immunostain always describes a whole set of neurones, criteria are given to determine the degree of structural variations within one neural set in a single individuum. Furthermore, this set of neurones establishes position markers that express the respective antigen for the different segments originating from a specific neuroblast. Therefore, the fate of a whole neural network that incorporates immunoreactive cells can be determined and it may be possible to find homologous structures throughout the different branches of the arthropod system (Weiss and Kupfermann, 1976). 3.1.4.4.1 Proctolin. Proctolin is one of the most intensively studied invertebrate neuropeptide (O’Shea, 1982). It was one of the first neurotransmitters to be used for the characterization of identified neurones (Rowell, 1976). Originally it was isolated from the cockroach, Peripfaneta americana (Brown, 1975; Brown and Starratt, 1975), and later it was characterized as a pentapeptide H-Arg-Tyr-Leu-Pro-Thr-OH (Starratt and Brown, 1975). The distribution of proctolin and proctolin immunoreactive neurones is well established for insects (Brown, 1977; Veenstra et af., 1985; Nassel and O’Shea, 1987; Blechschmidt et af., 1988), and crustaceans (Schwarz et al., 1984; Siwicki et al., 1985; Marder et al., 1986; Siwicki and Bishop, 1986; Stangier et af., 1986). It has been described as a peripheral neurotransmitter acting on visceral and skeletal muscles (Piek and Mantel, 1977; O’Shea and Adams, 1981; O’Shea and Bishop, 1982; Adams and O’Shea, 1983; Witten and O’Shea, 1985; Lange et a f . , 1986) and as a central neurotransmitter or neuromodulator (Eckert et al., 1981; Bishop and O’Shea, 1982; Agricola et af., 1985; Keshishian and O’Shea, 1985a; Veenstra et al., 1985; Siwicki and Bishop, 1986; Nassel and O’Shea, 1987). Additionally, it seems to play a neurohormonal role as a cardioactive substance in insects (Miller, 1983; Konopinska et af., 1986, 1988; Nassel et al., 1989) and crustaceans (Schwarz et af., 1984; Stangier et a f . , 1986). The investigations of Keshishian and O’Shea (1985b) on the early


49

development of the grasshopper revealed that proctolin is already expressed during embryogenesis. In the beetle Tenebrio molitor, 10 thoracic and 12 abdominal immunoreactive neurones were described in each neuromere (Breidbach and Dircksen, 1989). These neurones are present in the larva and persist throughout metamorphosis without changes in their principal structural characteristics. The structure and distribution of some of these identified immunoreactive neurones in Tenebrio (Breidbach and Dircksen, 1989) are comparable to neurones described for the cockroach (Eckert et al., 1981; Bishop and O’Shea, 1982; Agricola et al., 1985), the locust (Keshishian and O’Shea, 1985a,b) and the blowfly (Nassel and O’Shea, 1987; compare with Table 3). A comparison of the general distribution of thoracic and abdominal neurones of Tenebrio with those of the cockroach (Bishop and O’Shea, 1982) reveals striking similarities, which are also apparent for the terminal ganglion of both species (Eckert et al., 1981; Agricola et al., 1985). Using anatomical criteria (location of somata, topology of axonal projections, projection sites of the primary dendrites) and identical immunoresponse, Breidbach and Dircksen (1989) proposed homology of some of the identified neurones in the beetle with those of the blowfly, the locust and the cockroach (Table 3). TABLE 3 Comparison of proctolin immunoreactive neurons in the CNS of different insect species Tenebrio molitof Periplaneta americanab Periplaneta americana‘ Periplaneta americanad Schbtocerca niterne Periplaneta americand Calliphora erythrocephalag

PFN

AN1 type4 “1” type4

AN2

LMDN

“1”

LVCL type9 VM‘

MDN PNl typezh LDM/PC “3” t~pe2~ D sJ T1/2mv

This table compares the appropriate protolin immunoreactive VNC neurones for different insects and the terminology applied by the investigators. a Breidbach and Dircksen (1989); Eckert et al. (1981); Bishop and O’Shea (1982); Agricola et al. (1985); Keshishian and O’Shea (1985a); Witten and O’Shea (1985); Nassel and O’Shea (1987); only one of this neurone group is present; ’ for Tenebrio a descending projection was identified; j the Dsneurone belongs to a population that appears to be topologically equivalent to PN,of Tenebrio.

3.1.4.4.2 FMRFamide. FMRFamide-like immunoreactivity characterizes a family of neuropeptides composed of 7-18 amino acids each ending with a Arg-Phe-amide segment (for review, see Kingan et al., 1990). Antisera used for analysis of FMRFamide immunoreactivity characterize a whole class of related neuropeptides (Price and Greenberg, 1989). Immunohistochemical and physiological findings suggest that FMRFamiderelated peptides are neurotransmitters or neuromodulators in the CNS as well as neuromodulators or neurohormones at peripheral sites (Walther et al., 1984; Evans and Myers, 1986; White et al., 1986; Walther and Schiebe,

50


1987; Jenkins et a f . , 1989; Robb et al., 1989; Lundquist and Nassel, 1990). With FMRFamide immunohistochemistry the problem arises that several of the known antisera also recognize pancreatic polypeptides and crossreact with gastrin and the CCK in vertebrates (Price and Greenberg, 1989; Sossin et a f . , 1989). Thus, for credible information it is required to reveal the specificity of the antisera used. FMRFamide-like immunoreactivity has been employed to demonstrate homologous neurones in the VNC of different insect species (Veenstra, 1984; Veenstra and Schooneveld, 1984). Putative homologous clusters of FMRFamide-like immunoreactive neurones are present in the suboesophageal ganglia of the beetle Leptinotarsa lineata and the locust Locusta migratoria (Veenstra, 1984). Homologies at the level of identified neurones can be postulated by comparing FMRFamide immunoreactive neurones in the VNC of adephagan and polyphagan beetles (Breidbach, unpublished). Structural characteristics of a set of six bilaterally symmetric serial homologous neurones with somata clustered in the VNC ganglia have been identified in all species analysed so far, with only slight differences in structural details. FMRFamide-like neurones have been characterized previously in the adult brains from many insect species, including the lepidopteran Manduca sexta (Homberg et a f . , 1990, 1991), the honeybee Apis meffifera (Schurmann and Erber, 1990; Eichmuller et al., 1991) and the dipteran Drosophifa melanogaster (White et a f . , 1986). There is some evidence that most of these immunoreactive neurones are central interneurones partly associated with sensory neuropiles, or neurosecretory cells. During metamorphosis, both types of immunopositive neurones undergo significant changes in their appropriate peripheral projections (Hildebrand, 1985; Copenhaver and Truman, 1986). For the brain, it is difficult to attempt to homologize immunopositive cells. Various antisera have been used in descriptions of insect brain and VNC FMRFamide-like immunoreactive neurones (Eichmiiller et al., 1991; Schooneveld et al., 1992). It is also observed that the structural patterns of immunoreactive neurones vary in the different species. A comparison of the immunopositive neurones in the adult brain of Manduca and Tenebrio does not suggest immediate correlations of certain cell types (Homberg et a f . , 1990; Breidbach and Wegerhoff, 1993). The principal organization of FMRFamide-like immunoreactive neurones demonstrates striking differences in comparable neuropile areas of these two species: in the antenna1 lobe of Manduca about 80 prominent immunoreactive neurones were found, whereas in Tenebrio only three prominent immunoreactive neurones were located (Breidbach and Wegerhoff, 1993). Furthermore, in Manduca (Homberg et a f ., 1990), the protocerebral bridge shows massive immunostaining, which has no correspondence in Tenebrio (Wegerhoff and Breidbach, 1992). Comparison of Tenebrio (Breidbach and Wegerhoff, unpublished observations) with Apis meffifera (employing the same antiserum in both studies) shows conformity in the staining pattern of


51

Kenyon cells (Schurmann and Erber, 1990; Eichmiiller et al., 1991). Both species, Apis and Tenebrio, possess strata of immunoreactive structures that extend through the entire mushroom body neuropile. In both species, immunoreactivity of the Kenyon cells is restricted to axons and collaterals. A comparison of the brain of Tenebrio with that of the adult Drosophila shows structural conformities (White et al., 1986). In Drosophila, subpopulations of neurones are present. With respect to position of the clusters and the number of the neurones encountered in each cluster a correlation with immunopositive midbrain clusters in Tenebrio is suggested. With regard to the position and the number of the respective neurones, the Drosophila cluster SPI is comparable with cluster 2 in Tenebrio (Breidbach and Wegerhoff, unpublished observations) ; Drosophila cluster SP3 may correspond with cluster 1 in Tenebrio; size and position of Drosophila cluster LP2 (White et al., 1986) correspond with the cluster 8 of Tenebrio. However, morphological descriptions of the FMRFamide-like immunoreactive cells in Drosophila are not yet sufficiently detailed to homologize the neurones of both species. In Drosophila, analysis of the development of FMRFamide-like immunoreactive neurones using immunohistochemistry (White et al., 1986) and in situ hybridization (O’Brian et al., 1991; Schneider et al., 1991) demonstrate the persistence of the principal features from the late embryo to the adult. Obviously, all larval immunoreactive brain neurones persist to the adult stage, showing an immunoreactivity throughout all stages of pupal development (White et al., 1986). The latter is confirmed by analysis of the expression pattern of the FMRFamide neuropeptide gene during metamorphosis of Drosophila (O’Brian et al., 1991). These results are comparable with those found for the development of FMRFamide-like immunoreactivity in Tenebrio (Urbach et al., 1993). The expression of the FMRFamide gene in the embryo of Drosophila (immunohistological detection: from stage 15 onwards; gene expression from stage 16 onwards; Schneider et al., 1991) is comparatively late, compared to Tenebrio (immunopositive brain neurones in the 60% embryo). 3.1.4.4.3 Crustacean cardioactive peptide (CCAP). One of the most interesting neuropeptides that may be employed for a comparative analysis in Arthropoda seems to be the crustacean cardioactive peptide (CCAP). Originally, it was described as a cardioacceleratory substance stemming from extracts of the pericardial organs of the shore crab, Carcinus maenas. It was biochemically characterized by Stangier et al. (1987) and represents a cyclic nona-peptide (PFCNAFGTGC-NH2). Radioimmunological studies demonstrated the occurrence of CCAP immunoreactivity in all ganglia of the CNS of Locusta migratoria (Stangier et al., 1989). Evidence was provided for the presence of endogenous CCAP in an insect CNS by sequencing the peptide from purified CNS extracts. CCAP immunoreactive interneurones and neurones with projections to visceral muscles and to neurohaemal areas in the thorax and abdomen were discovered in the locust (Dircksen et al.,

52


1991) and the meal beetle (Breidbach and Dircksen, 1991). Thus, structural and topological similarities of CCAP immunoreactive neurones of the VNC of the CNS of the crab and the locust are apparent. Therefore, CCAP seems to be most promising for a comparison between different Mandibulata. Since CCAP immunoreactivity is also expressed in the nervous system of spiders (Breidbach, 1992a), a comparison can be extended to Arthropoda in general. CCAP is expressed early in a 45% embryo of Tenebrio (Urbach et al., 1993), which may allow a description of the cell lineage of the respective neuropeptide expressing cells. The analysis of CCAP immunoreactivity in Locusfa reveals a set of identified bilaterally symmetrical neurones in all VNC ganglia (Dircksen et al., 1991). Structural and topological comparison of these neurones in the locust and the NT1- and NTz-neurones in Tenebrio molitor suggests an interspecific homology of these neurones (Fig. 8). Position of somata, central pathways and contralateral projections of these neurones are essentially similar (Fig. 9) (Breidbach and Dircksen, 1991). With regard to putative neurohaemal release sites a different situation occurs. In Tenebrio these sites are formed by both NTl and NT2 (Breidbach and Dircksen, 1991). For the putative homologous type 2 neurone in Locusfa similar projections to these peripheral sites are missing (Dircksen ef al., 1991). The neurohaemal sites of the locust are supplied by type 1 neurones and an ipsilaterally projecting type 4 neurone, for which a counterpart seems to be missing in the CNS of Tenebrio (terminology after Dircksen et al., 1991). Similarly, there is no obvious correspondence between the distribution of brain interneurones in Tenebrio and Locusta. In Tenebrio, 10 protocerebral immunoreactive neurones are present, whereas in the locust more than 100 neurones are seen distributed throughout almost all parts of the brain (Dircksen and Homberg, unpublished). Preliminary experiments on the localization of CCAP immunoreactive neurons in the larval CNS of Calliphora vicina and Drosophila melanogaster reveal putative serially homologous sets of contralaterally projecting interneurones in the fused VNC neuromeres. This situation is topologically identical to the position of the NTl- and NTz-neurones in Tenebrio (Dircksen et al., unpublished). Furthermore, the number, structure and location of immunopositive brain interneurones resemble those of Tenebrio (Fig. 10). Hereby, Drosophila expresses only six immunoreactive neurones. Projections and arborization patterns match with those of a first instar larva of Tenebrio, which, incidently, also has only six bilaterally symmetrical immunopositive neurones. Apparently, the set of CCAP immunoreactive neurones, described for Tenebrio (Breidbach and Dircksen, 1991), represents a set of neurones whose structures are phylogenetically highly conserved within Holometabola. Furthermore, these neurones represent a group of neurones, for which the established criteria of homology are also applicable with respect to several locust neurones (Fig. 9).


53

FIG. 8 Structurally similar pattern of projections of CCAP immunoreactive neurones in the mesothoracic ganglion of (A)adult Locusta migruforia (scale bar 100 pm) and (B) Tenebrio molitor (scale bar 20 pm). Note similarities in pathway characteristics of neurone 1 and 2 (1, 2; white triangles). (From Dircksen et al. (1991) (A) and Breidbach and Dircksen (1991) (B).)

A

FIG. 9 Homologous individual CCAP immunoreactive neurone (type 2) in the mesothoracic ganglion of (A)Locustu migrutoriu (scale bar 100 pm) and (B) of larval Tenebrio molitor (scale bar 20 pm). Note correspondence in branching characteristics (triangles and arrow, respectively). a, anterior; p, posterior. (From Dircksen et ul. (1991) (A)and Breidbach and Dircksen (1991) (B).)

54


A

FIG. 10 Homologous CCAP immunoreactive neurones in the brain of holometabolous insects; frontal views, wholemount preparations, camera lucida drawings. (A) Drosophila melanogaster, 3rd larval instar. (B) Tenebrio molitor, 1st larval instar. Note presence of three immunoreactive somata (black triangle) in each hemibrain, with similar pathway (arrows) and branching characteristics. A , ascending projections of VNC immunoreactive neurones; D , dorsal; V, ventral. Scale bar 50 pm. (From Dircksen and Breidbach (unpublished) (A) and Urbach et al. (unpublished) (B1.1

Interestingly, this set of neurones bears some structural pecularities, which are also found in serial homologous crustacean CCAP immunoreactive neurones (Dircksen and Keller, 1988). To decide whether such a phylogenetical similarity is likewise seen at the single cell level, it is necessary to describe the cell lineages of the CCAP immunoreactive neurones in both groups of Mandibulata. 3.1.4.4.4 Serotonin. In a review on the physiological roles and distribution of serotonin in the insect nervous system Nassel (1987b) suggests that neurones exhibiting this immunoreactivity constitute part of the basic “Bauplan” of insects (Bishop and O’Shea, 1983; Tyrer et af., 1984; Homberg and Hildebrand, 1989). In the VNC, serotonin immunoreactive neurones form a series of serially homologous neurones (Figs 6 and 7), similarly expressed within the nervous systems of Orthoptera, Diptera and Coleoptera (Taghert and Goodman, 1984; Tyrer et af., 1984; Hustert and Topel, 1986; Rheder et af., 1987; Breidbach 1987a,b; Cantera and Nassel, 1987; Valles and White, 1988). In the grasshopper these serially homologous neurones constitute a group of four bilaterally symmetric cells, their somata lie in the posterior ventro-lateral neuromers of thoracic and abdominal ganglia. The two neurones per hemiganglion constitute two neurone types,


55

both form contralateral projections and exhibit an ascending and a descending axon. The immunoreactive cell of type S1 (Taghert and Goodman, 1984) forms a brush-shaped arborization in the anterior half of the contralateral neuropile of the ganglion. Axons of both neurones form a fascicle. The cell lineage of these cells is established. Both derive from the same precursor. Their development can be followed up. Serotonin expression was found in the neurones commencing at about 55-58% of embryogenesis. This is of interest because this phenomenon occurs before completion of the morphological development of these cells (Taghert and Goodman, 1984). A structurally similar distribution of serotonin immunoreactive cells within the VNC is also apparent in the dipterans Drosophila melanogaster (White and Valles, 1983; Valles and White, 1988), Calliphora erythrocephala, and Sarcophaga bullatta (Nassel and Cantera, 1985; Cantera and Nassel, 1987), in the beetle Tenebrio molitor (Breidbach, 1987a,b, 1991), in the cockroach (Bishop and O’Shea, 1983) and the dragonfly (Longley and Longley , 1986). All this suggests interspecific homology. Taghert and Goodman (1984) described a third immunoreactive neurone in the hemineuromers of the locust prothoracic and suboesophageal ganglion. A serotonin immunoreactive cell with a soma at a similar position was also found in Calliphora (Nassel and Cantera, 1985) and in Tenebrio (Breidbach, 1987b, 1991). Thus, within the Insecta the serotonin immunoreactive neurones seem to constitute part of the basic “Bauplan” of the insect VNC. In crustacean species, lobster (Beltz and Kravitz, 1983), Procambarus clarkii (Real and Czternasty, 1990) and Oniscus asellus (Breidbach et al., 1990), VNC serotonin immunoreactive cells are also present which are similar to those of insects both in number and principal topology. Therefore, serotonin immunoreactive cells constitute further candidates for a cell population which might be homologous in Mandibulata. For the insect brain, analysis of serotonin immunoreactive neurones outlines some basic similarities between insect species. Such neurones were identified in the brain of the hemimetabolan Periplaneta americana (Bishop and O’Shea, 1983; Klemm et al., 1984), Locusta migratoria (Klemm and Sundler, 1983; Konings et al., 1988b), Schistocerca gregaria and S . americana (Homberg, 1991), Rhodnius prolixus (Lange et al., 1988) and of the holometabolan Drosophila melanogaster (White and Valles, 1983; Valles and White, 1988), Calliphora erythrocephala, Sarcophaga bullata (Nassel and Cantera, 1985; Nassel et al., 1987), Apis mellifera (Schurmann and Klemm, 1984; Rheder et al., 1987), Manduca sexta (Granger et al., 1989; Homberg and Hildebrand, 1989), and Tenebrio molitor (Breidbach, 1987d, 1990b). However, only a few studies (Table 4) allow a comparison of cellular structures in the insect brain neuropile. Such an analysis of identified elements is essential to establish possible homologies of serotonin immunoreactive neurones. Klemm et al. (1984) described the main pathways of

56


serotonin-immunoreactive neurones in the brain of the cockroach, which match with those of Tenebrio. Presently, only for Tenebrio and Manduca are the anatomical descriptions detailed enough to strive for homologization of protocerebral immunoreactive neurones. For the deutocerebrum homologizations may be possible for both holo- and hemimetabolan species. The serotonin immunoreactive neurone of type CPNl in Tenebrio (Breidbach, 1990b) probably is homologous to a similar neurone in Manduca (Kent et a f . , 1987; Granger et a f . , 1989) and in the hemimetabolan Periplaneta americana (Salecker and Distler, 1990). Recent studies of the metamorphic development of Tenebrio indicate that this neurone persists throughout metamorphosis without major alterations of its arborization pattern (Breidbach, 1990b; Wegerhoff and Breidbach, 1992). TABLE 4 Identified serotonin neurons in the holometabolan central hemi-brain Drosophila melanogastef (larva) Calliphora erythrocephalab (larva) Sarcophaga bullata" (larva) Apis melliferad Manduca sextae (larva) Manduca sextd (adult) Tenebrio molitor': (larva) Tenebrio molitof' (adult)

11 15 16 20 18-20 20-21' 22 22'

Number of serotonin immunoreactive neurones in the brain, as identified by several investigations. a White and Valles (1983); b s c Nassel and Cantera (1985); Schurmann and Klemm (1984); Granger ef al. (1989); Homberg and Hildebrand (1989); g*h Breidbach (1990b); neurones in the optic neuropile were not considered.

Comparison of larval Tenebrio with larval Manduca strengthens the assumption that serotonin immunoreactivity marks a phylogenetically highly conserved set of neurones. For Manduca, a neurone passes through the commissure two (Granger et al., 1989) which corresponds to a neurone, which crosses via the deutocerebral commissure in Tenebrio projecting to the contralateral accessory lobe (Breidbach, 1990b). Neurones which project to the posterior-ventral protocerebrum with projection sites in the superior protocerebrum are present in both insects. Only for the dorso-lateral protocerebral neurones is a variation seen. For Tenebrio three dorso-lateral protocerebral neurones were identified, whereas for Manduca only one cell was labelled. The tritocerebral ascending neurones characterized in Tenebrio (Breidbach, 1990b) were not described in Manduca larva (Granger et al., 1989). Homberg and Hildebrand (1989) display detailed reconstructions on the single cell level for adult Manduca. Together with the results mentioned in the present study, the idea is supportive of a principally similar organization of serotonin immunoreactive neurones manifested in different insect species. The cells that reach the central body are apparent in both species, similar in topological representation, principal structure and number


57

(Wegerhoff and Breidbach, 1992). Contralaterally projecting neurones passing through the deutocerebral or fronto-median commissures are equivalent in both, Manduca and Tenebrio. The ipsilateral projecting lateral protocerebal neurone of Manduca (Homberg and Hildebrand, 1989) corresponds topologically and structurally with the neurones of cluster 1 in Tenebrio (Breidbach, 1990b). However, the arborization patterns of single neurones vary in the different species. So far, for the brain structural conformity is less pronounced than for serotonin immunoreactive interneurones of the VNC. Though, at least for those neurones which correspond individually in their topological and structural organization, homology can be suggested. Such a phylogenetic conservation in structural organization of identified immunoreactive brain interneurones is characterized up to the level of fine arborization patterns, for different beetles. Also for specific brain neuropile areas, such as the mushroom bodies, identified serotonin immunoreactive neurones exhibit structural similarities in their fine arborization patterns (Breidbach, 1990~). Thus, it is suggested that within the VNC and the brain of different insect species, neurones can be determined to represent homologues. All these homologues constitute a complex network of phylogenetically conserved structures. They seem to form a small interconnected network, partly conserved even in its finer meshworks. Interestingly, such phylogenetically conserved neurones are also found within sensory neuropiles, which is apparent for the serotonin-immunoreactive deutocerebral neurones. 3.2

MYRIAPODA

While it is generally accepted that the Insecta are of monophyletic origin, this assumption is not yet fully established for the Myriapoda, which are split up into Diplopoda and Chilopoda. Even though, Insecta and Myriapoda represent sister-groups comprising the Tracheata (Gupta, 1979). Both groups show many features of morphology and functional anatomy in common (Manton, 1972). Also for early embryology common traits are observed (Anderson, 1973). Embryological analysis has outlined that, in principle, neurogenesis in myriapods is comparable with that in insects (Dohle, 1964, 1974; Knoll, 1974; Hertzel, 1984; Whitington et al., 1991). This general comformity was recently supported by an analysis of molecular probes (Field et al., 1988), though the phylogenetic divergence of both groups may be greater than previously assumed. However, differences in the expression of engrailed proteins and in the origin of pioneering axons in the CNS are apparent (Whitington et al., 1991). Peculiarities such as early separation of the left and right sides of the ganglionic primordia by extraembryonic ectoderm was found in the studied species (Ethmostigmus rubrips). This might be explained as an early deviation from the general

58


plan of embryogenesis otherwise observed in the group of Tracheata (Korschelt, 1936). Therefore, it is open as to whether such differences vary from the fundamentally conservative mode of tracheate development. The supposed close association with the insects and the rather homogenous segmentation should have stimulated comparative studies. However, such studies for Myriapoda are scarce. They should be directed towards an understanding of the “basic segment”. This statement is valid also with respect to the nervous system (see the appropriate short chapter in Bullock and Horridge (1965); for brain structures, see Joly and Descamps (1987)). As has been mentioned, studies on chilopodan neurogenesis show interesting differences between insects and centipedes (Hertzel, 1984; Whitington et al., 1991). For some comparative aspects of early neurogenesis in Diplopoda and Chilopoda with other Arthropoda groups and Annelida and Onychophora, see Muiioz-Ceuvas and Coineau (1987). In insects, the CNS originates from a neuroblast array segmentally reiterated along the early embryo (Thomas et al., 1984). However, in Chilopoda neurogenesis and ganglion formation occur independently of the generation of a typical neuroblast array. As mentioned above, an antibody raised against the engruiled protein repeatedly stains rows of cells indicating the posterior rim of the segments in Insecta and crayfish (Patel et al., 1989a,b). The engrailed protein is also expressed in the early centipede. Though the marked cells differ in their location, there is no antibody binding to the ganglionic primordia. Which are the consequences of such differences? Is there any reason to search for possible homologies in the neuromuscular system of Insecta and Myriapoda? Is there any reason to suppose that apparent “homologous” muscles are innervated by apparent “homologous” MNs stemming from different neural lines? 3.2.1 Motoneurones A general introduction to the nervous system of Chilopoda is presented by Fahlander (1938). Lorenzo (1960) describes the cephalic nervous system of a more primitive centipede (Arenophilus, Geophilomorpha), while Rilling (1960, 1968) gives a detailed description of the neuromuscular system of a more advanced centipede (Lithobius, Lithobiomorpha). Even though there is some heteronomy distinguishing adjacent segments in Lithobius, muscles and nerves exhibit a rather uniform design in view of a serial homology. Recently, the MNs have been considered which supply the DLMs (Heckmann and Kutsch, unpublished). We have chosen this muscle system, since an equivalent system is also apparent in Insecta and Decapoda. As also observed for both latter arthropod groups, in Chilopoda, the relevant MNs are distributed between two ganglia. In Lithobius, there are 24 MNs in the next anterior ganglion and 4 MNs in the ganglion immediately associated with the position of the muscle system. When compared to insects it


59

becomes apparent that in Lithobius the complctc set of DLMs is composed of more individual muscles, consisting of muscles spanning from uni- to trisegmental spaces. A selective stain employing only the unisegmental muscles results in a considerable reduction of the stained MNs (Fig. 11; see also Table 2). Even so, the morphology of these MNs is rather different when compared to the MNs serving an apparently equivalent muscle in insects. Therefore, we tend not to accept an immediate homology of the motor system of both groups. With respect to the apparent similarities based on the major split of the MNs into two ganglia and their position, it still remains obscure how different neurogenesis strategies (see above) will result in such resemblance. 3.2.2 Interneurones and sensory neurones Apart from some gross histological characterizations of myriapod neuroendocrine cells (Sahli, 1974; Joly, 1979), we are not aware of a recent description of non-motoneural VNC or brain neurones in Myriapoda. Recently one of us (Breidbach, unpublished) has started to map immunoreactive neurones (FMRFamide-like, CCAP-like and serotonin-like immunoreactive neurones; Fig. 12). Preliminary results show serial homologues; for serotonin-like as well as CCAP-like interneurones common structural features are seen (position and number of somata, topology of primary branches) with certain neurones in insect VNC neuromers. However, before engaging in discussions of possible homologies of neurones for Myriapoda and Insecta more detailed studies of cell lineages are required (Dohle, 1964, 1974; Knoll, 1974; Hertzel, 1984). Also, detailed neurohistology is still meagre for myriapods. It may be of interest that data concerning the brain morphology led Bullock and Horridge (1965) to state “The brain structure of Myriapoda is not particularly helpful in showing their relationships to other arthropod groups, and, in particular, provides no evidence that they are related to the ancestors of the insects”. With respect to receptor organs there is some more information available which may be compared with other Arthropoda groups (Wright, 1976). For Insecta and Crustacea, muscle stretch receptors are present in the thorax and abdomen. Such receptor organs were also discovered in centipedes, unior bisegmentally arranged (Rilling, 1960; Osborne, 1961, cited in Finlayson, 1976; Varma, 1972). Bipolar and multipolar neurones are also present. Generally, for centipedes several nerve cells are attached to each receptor muscle or an associated connective tissue band, whereas for insects usually only one neurone is present (Finlayson, 1976). Similar muscle receptor organs apparently exist also in the coxa-trochanter joint of each leg of Lithobius (Rilling, 1960). Similar organs are known in Crustacea, and

60

W. KUTSCH AND 0. BREIDBACH ant

G2

Q3

MScdh DLMB Tei

FIG. 11 Camera lucida drawing of the neural set innervating unisegmental DLMs of the centipede Lithobius. In this preparation a contralateral soma and two midline cells in the anterior ganglion are missing. For this preparation, the nerve to the DLMs of the 3rd segment has been stained resulting in the typical separation into two neural groups: anterior ganglion with ipsilateral somata and posterior group with contralateral somata. G2, G3, ganglion of the 2nd and 3rd leg-bearing segment; N 1-8, lateral nerve roots; ant, anterior; post, posterior. (Modified after Heckmann and Kutsch (unpublished).)


61

FIG. 12 Two groups of serial homologous interneurones (characterized by CCAP immunoreactivity) in two VNC ganglia (S8 and T, the terminal ganglion) of the chilopod Lithobius fortificnns. Note presence of six bilateral symmetric central descending neurones (CDN) and two descending neurones (DN) per hemiganglion; note correspondence indicated by equal arrows and triangle. Scale bar 50 pm. (From Breidbach (unpublished).)


62

appear in rare cases in Insecta (Markl, 1966). For insects usually joint movements or cuticle stretches are monitored by chordotonal organs. Such typical receptor structures seem to be missing in Chilopoda. Distortion of the membranes seems to be measured by receptor neurones attached to the epidermis via free sensory endings (Rilling, 1960). Externally, apparently mechanoreceptive hairs are present which often are united to segmentally repeated hair fields (Rilling, 1960). Although the peripheral sensory array is elaborated, the paucity of different sensory types and structures is remarkable for Chilopoda. Also in this respect the difference should be mentioned in comparison with Crustacea and Insecta. This difference almost excludes an approach to search for possible homologies among these arthropodan groups.

3.3

CRUSTACEA

There is a wealth of information available on the general structure of the Crustacean nervous system, and on a comparison of its structure among the different species and orders (e.g. Bullock and Horridge, 1965; Atwood, 1982; Sandeman, 1982; Nassel and Elofsson, 1987). Although similarities usually are apparent, homology is not necessarily implied. For the crayfish Procambarus clarkii a first rather accurate count of neurone numbers has been given and single identifiable neurones have been described (Wiersma, 1957). Information on the general structure of selected ganglia is available, especially for the more advanced Decapoda (Retzius, 1890; Kendig, 1967; Tsvileneva et al., 1976; Skinner, 1985a,b; Chaudonneret, 1985; Kondoh and Hisada, 1986). Early work on embryonic lobster demonstrated reiterated individual nerve elements (Allen, 1894, 1896). All studies not only point to serial homology but equivalences seem to occur throughout the Decapoda and they might even be characteristic for all Crustacea. 3.3.1 Motoneurones The improvement of selective stains has allowed the identification of individual MNs, usually combined with physiological studies (Wine et al., 1974; Wilson and Sherman, 1975; Wiens, 1976; Mittenthal and Wine, 1978; Nagayama et al., 1983; Mercier et al., 1991; Wiens and Wolf, 1993; Fig. 13). It becomes apparent that each MN has its specific soma, size, position and structure, repeated from preparation to preparation with only a small variability in details (e.g. Wiens, 1976). Functional antagonists (crayfish claw opener vs. closer) are structurally different, whilst MNs serving synergistic muscles are markedly similar. The question of serial homology has been raised previously by comparing groups of MNs along the crayfish abdominal nerve cord (Mittenthal and Wine, 1978). A group of three clusters (flexor


T3

63

/

FIG. 13 Camera lucida drawing of the neural set innervating the thoracic DLMs in a Crustacea (Zdotea balthica: Isopoda). Owing to a stain of the DLMs of the fourth segment, MNs are seen in the fourth (T4) (with contralateral somata) and third ganglion (T3) (with ipsi- or medial-positioned somata); one soma is even detected in the second thoracic ganglion (T2). N 1-3, segmental nerve roots, anterior is up. (From Schneider (unpublished).)

64


MNs) can be described for each of the five anterior abdominal segments. For certain ganglia (abdominal ganglia G2-G4), the motoneurons are more similar while for others (Gl+G5) differences of the choosen motoneurons are apparent (Mittenthal and Wine, 1978). Such a detailed analysis has prompted a comparison with other Decapoda (as well as with Insecta, see below). The grouping and structure of MNs suggests a homology of these elements among the Decapoda. In a very detailed study, Wiens and Wolf (1993) have compared the three inhibitory MNs supplying the thoracic limbs of three crayfish species employing differential backfills, Lucifer yellow intracellular fills, immunohistochemistry and electrophysiology (see Fig. 14A). Whether studying the chelipeds or the walking legs, for these specific MNs there are so many details in common that a definitive example is postulated, not only of serial homology, but also of homology among closely related species (for a comparison with Locusta, see below). In a correlation of function and structure, it is suggested that the opener inhibitor is more similar to its functional synergist (closer excitor) than to the other two inhibitors. Whether this similarity is required by constraints of common premotor elements to synergists (Wiens and Gerstein, 1975, 1976) needs further study; for Schistocerca, a similar argument that synergistic (flight) MNs are more similar amongst each other than compared with their antagonists, has been discarded (Siegler et al., 1991). Little information exists on the general structure of the NS in nonMalacostraca and a detailed description of identified neurones is still missing. An extended analysis of Crustacea comprising a group of such a wide range of phylogenetic divergence (Bowman and Abele, 1982) could be rewarding in pursuing the concept of possible homologies throughout evolution. In recent studies the distribution of certain immunoreactivities, such as serotonin, proctolin (Beltz et al., 1990) or neurohormones (Mangerich et al., 1986) was investigated. Serial repetition is observed from the thoracic to the terminal abdominal ganglia. Individual reiterated elements can be detected and homologies among species are apparent. Certain time windows are apparent for embryonic and larval ontogeny indicating that relevant studies also require a consideration of the developmental stage (for several references, see Beltz and Kravitz, 1983; Siwicki and Bishop, 1986; Siwicki et al., 1987; Real and Czternasty, 1990). An interesting case is the comparison of MNs in a system manifesting bilateral asymmetry. For example, claw asymmetry occurs among Crustacea. Such an asymmetry is seen, in parallel, not only in the form of the claws and the muscle mass but also in soma size, dendritic fields of the MNs and the number of receptor neurones (Mellon, 1981; Young and Govind, 1983; Govind and Pearce, 1985). While in larval and early juvenile stages of Hornarus symmetry of both sides occurs, some asymmetry develops for the


65

crusher and cutter during later juvenile and adult stages (Govind, 1984). Hereby, the original symmetric innervation becomes unequal, involving not only a change in the neural supply but also a change in the activity of the appropriate MNs. Structural and functional switches can also occur in later stages of a shrimp (Alpheus heterochelis) after manipulation of the peripheral nerve (Mellon, 1981). Whatever the causative factor is, it becomes apparent that a selective influence on neural structures and their properties becomes possible by external cues. Such results should be taken into consideration when inspecting regenerating neural structures. At the tip of all five pairs of thoracic limbs of Homarus a chela is present, differently organized (from claw to a small appendage) throughout the segments. Each of the chelae are operated by an opener and closer muscle. Whilst the opener muscle is rather similar for all serially homologous limbs, muscle fibre correlation (fast vs. slow) changes for the closer muscle (Mearow and Govind, 1986). With respect to the number and type of MNs there is, in general, a similarity in the topology and morphology of these MNs with only slight differences concerning the branching pattern or distribution of fast axon synapses. This indicates that differences in the periphery need not be paralleled by changes in the basic neural circuit; although this notion does not imply that the activity of the different elements is equal. A complete set of serial homologous MNs has been postulated to serve the abdominal swimmerets in Homarus. Apparently, the different motor axons grow in a common group in the embryo, even before the external appendages are formed and only later do the axons disperse and rhythmic activity commences (Kirk and Govind, 1992). Immunohistochemical studies also reveal apparent serial homologues in Crustacea. However, often this technique does not resolve cellular structures in great detail, which might result in some uncertainties of equivalence. Mercier et al. (1991) show catecholaminergic neurones in the abdominal ganglia of Procambarus. Equivalent positions are apparent for several smaller somata in different segments and for two larger cells in two of the segments with similar dendritic fields and peripheral target. 3.3.2 Receptors There is a wealth of information concerning receptor systems in Crustacea (for summary: Bullock and Horridge, 1965; Mill, 1976a,b; Laverack, 1987). Parallel to the point raised above for the MNs, receptor systems have been studied in more detail in the Decapoda than in any other Crustacea group; studies of the latter would add valuable information to questions concerning homology and phylogeny. Very often, receptor organs are placed superficially. In this location, they may respond to external influences, such as medium currents, or they may

66


be used for proprioception, such as in the cases of placement in the limb joint or between segment borders. There are rows of cuticular pegs (CAP-organ) (Alexandrowicz, 1972) which are repeated at equivalent joints throughout all the thoracic legs of Hornarus and they are also seen along the appendages in the mouth region (Laverack, 1976; Wales, 1976). These receptors are present in early larval stages, but increase in numbers of sensilla at each moult. Homologous receptor fields are formed in a variety of Decapoda at corresponding positions (Laverack, 1976), though in Brachyura they are totally absent. For the latter group, embryological studies should reveal whether these organs are present early in ontogenesis, but are lost during further development. The major internal proprioceptors in Crustacea are the chordotonal organs and muscle receptor organs. After its initial description by Alexandrowicz (1951) for the lobster, this ensemble of sense cells attached to dorsal muscle fibres has been studied in much detail. The tonic and phasic receptor cells are observed in all abdominal and thoracic segments. Physiological and histological data show homologous receptors in almost all Decapoda, except for Cancer where it seems to.be missing (Pilgrim, 1960). In addition, a similar proprioceptor, the muscle receptor organ, is present at the body-coxa joint. The appropriate receptor cell somata are positioned within the ganglion (Alexandrowicz and Whitear, 1957; Bush, 1976), which mirrors an exceptional receptor type in the approximate leg joint of locusts. As a further characteristic these crustacean receptor cells do not propagate active spikes along their peripheral processes. This peculiar receptor organ is present all along the body, from the 2nd maxilla to the uropods, in a variety of Decapoda (Laverack, 1987). The chordotonal organs represent another type of mechanoreceptor. They have a variety of configurations and positions in the body; however, for our purpose it is of importance to state that similar receptor ensembles are present (with some exceptions) all along the body segments, appendages and in different Decapoda (Mill, 1976b; Laverack, 1987). An observed apparent deviation from serial homology has even been used as an argument that the paragnath, a small bilaterally symmetrical structure behind the mouth, characterizes an extension of the mouth region, but does not represent a true appendage (Laverack, 1987). As an interesting case of bilateral asymmetry the development of the two dimorphic chelae of the first thoracic leg has been mentioned above. The motor system is rather equal, though there is some minor difference in MN size and muscle control. However, for the sensory system a clear lateralization in number and size of afferent axons is discerned (Young and Govind, 1983; Govind and Pearce, 1985) with a factor of up to 1:4 favouring the larger claw. The target of these extra sensory fibres has not been determined. However, it becomes clear that peripheral receptor activities are involved in the transformation and maintenance of this system (Govind and Pearce, 1986).


67

There are plenty of hairs and hairfields distributed along the body and appendages. They serve as mechanoreceptive and chemoreceptive inputs (Bullock and Horridge, 1965; Laverack, 1987). It is expected that serial as well as interspecific homology also exists for these receptors but comparative approaches are needed to substantiate this assumption. 3.3.3

Interneurones

Rhythmic movements in Crustacea, whether employing the thoracic legs or abdominal appendages, appear to involve central motor programmes. Since such programmes in their basic forms are produced by isolated ganglia (or even hemiganglia; Hughes and Wiersma, 1960; Ikeda and Wiersma, 1964) it is surmised that not only MNs but also INS are serially reiterated. Stein (1971) documents the co-ordination of swimmeret MNs in crayfish by ipsilateral hemiganglionic oscillatory neurones successively coupled to each other by segmental co-ordinating fibres. Actually, some apparently homologous INS have been described (corollary discharge interneurones; Krasne and Wine, 1977); even so, homologous elements have been detected in only a few segments. There are four excellent classical studies demonstrating specific IN types which due to their serial repetition are supposed to be serial homologues (Allen, 1894, 1896; Bethe, 1897, 1898). The first pair of studies shows the potential of investigating an embryo with a clearly separated nerve cord in Hornarus. The second pair of studies shows the repetition within the fused complex of the thoracic ganglia of Curcinus, indicating the segmental neural sets. However, whether interspecific homologues can be identified awaits further comparative studies. The pioneering work of Wiersma and his colleagues demonstrated the existence of long intersegmental INS that either received inputs from specific sensory areas or transmitted information from higher brain centres to the lower centres of the ganglionic chain (e.g. circumoesophageal connective: Wiersma, 1958; abdominal connective: Wiersma and Hughes, 1961). Such excitatory or inhibitory command elements (for the dispute on the term “command”, see Davis and Kovac, 1981; Davis, 1985) can be associated with certain behavioural acts (Evoy and Ayers, 1982; Wine and Krasne, 1982). All these studies are mainly restricted to lobster and crayfish. It is to be expected that similar circuits with similar through-fibre systems exist, at least in the group of Decapoda. Comparative studies should yield a wealth of information on interspecific homologues on the basis of identified INS, identified morphologically as well as physiologically (e.g. substance-P-like neurons in the brain of crab and crayfish, Sandeman el al. 1990). Among the command neurones there are two prominent ones per side which, due to their size, can easily be identified as individuals: the medial and lateral giants (Wiersma, 1958; for their involvement in crayfish escape behaviour, see Wine and Krasne, 1982). Both giants appear to be well-

68


formed during embryogenesis (Allen, 1894, 1896), their persistence, growth and physiology has been followed up, from juvenile to the adult stage (Govind and Lang, 1967). The medial giant is a single cell with its soma positioned in the supraoesophageal ganglion. The lateral giant is composed of a series of segmentally reiterated single neurones, for each segment the soma is positioned laterally and the axons join each other via electrical synapses (Wiersma, 1947; Watanabe and Grundfest, 1961). This is an interesting system in which apparent serial homologous elements are coupled tightly to each other, eventually generating a single collective element. 3.3.4 Comparison of neural structures in Insecta and Crustacea Since both groups represent members of the Arthropoda, it is of interest to see whether analysis of Insecta and Crustacea can provide information on homologous structures. Such relevant comparative studies might reveal a basic “Bauplan” of a common arthropodan ancestor. With respect to neurogenesis, Thomas et al. (1984) showed that not only distantly related Insecta but also a Crustacea (the crayfish Procambarus) follow a similar programme of neurogenesis. This calls for a common embryonic plan “implying an early and subsequently conservative evolution of the developmental programme for the proto-insect (and perhaps protoarthropod) CNS”. Later, a sequential expression of segmental proteins (such as engrailed) during embryogenesis has been demonstrated for Insecta and crayfishes (Pate1 et al., 1989b). Skinner (1985a,b) describes the structure of an abdominal ganglion of Procambarus. With respect to existing studies of insect ganglionic architecture (e.g. Tyrer and Gregory, 1982; Watson and Pfliiger, 1987) there are many anatomical parallels. Skinner herself elaborates a nomenclature for several ganglionic structures adapted from knowledge on Insecta. This similarity in the basic plan of the ganglia of both groups suggests that homologous structures may be present even at the level of the individual neurone. At the level of identified neurones there are also several cases that point towards homology. Mittenthal and Wine (1978) have mentioned the similarity of somata with respect to position and central structures of MNs serving the dorsal abdominal flexors in Decapoda compared with those of the dorso-longitudinal muscle in a cricket. Parallel studies of an aquatic Isopoda (Idotea balthica) (Fig. 13) demonstrate several similarities with identified insect MNs, that suggest common phylogenetic traits (see also Table 2). However, there is one cell type (the DUM-cells) in Insecta for which a counterpart in Crustacea appears to be missing. Since DUM-cells proliferate from the median neuroblast it is not clear if this specific neuroblast is not present, or has changed its cell lineage, in Crustacea.


69

A very convincing example of possible homologous neurones in both Arthropoda groups is recently given by Wiens and Wolf (1993). There are several clusters of cell bodies exhibiting GABA-like immunoreactivity in the thoracic ganglia of Locustu and Orconectes. The distribution of these clusters propounds an arrangement not depending on mere accident but on common developmental processes. Locusts and crayfishes possess three inhibitory neurones which serve the leg muscles. The position of the cell bodies, the axonal branching and the pattern of muscle innervation are so similar that the authors associated each individual inhibitory neurone with its corresponding cell in the other group (Fig. 14). Some differences, such as an unusual axon routing, seem of no immediate advantage for evolution and there appears to be “no functional importance or selective value in itself”. Since they are “not likely (to be) the result of convergent evolution”, their similarity must be interpreted in terms of the “particular significance for the phylogenetic relationship of crustaceans and insects” (for some scattered information on similar immunoreactive elements in Insecta and Crustacea, see section 3.1.4). It is interesting to compare sensory structures among Insecta and Crustacea that exhibit similar morphological structures and ultrastructure (for proprioceptors, see Mill, 1976a). However, due to the diversity of the sense organs involved it becomes difficult to “prove apparent homologies”. It has been mentioned that the cell bodies of receptors in the coxal region of Crustacea are positioned in the thoracic ganglia (Alexandrowicz and Whitear, 1957; see also Bush, 1976). A similar unusual arrangement for mechanoreceptive neurones has been described for the coxal joint of the locust (Braunig, 1982b). This similarity suggests some sort of homology, even though a functional difference exists: while the insect organs produce action potentials, the crustacean receptors transmit their signals via graded potentials (Bush, 1967; Ripley et ul., 1968; however, for other mechanoreceptors with central somatas, “spiking” transmission has been shown: Pasztor and Bush, 1983, 1989). However, as yet, there are numerous dissimilarities that prohibit a clear-cut decision.

3.4

CHELICERATA

Based on the classical papers of Holmgren (1916), and Hanstrom (1928a), several brain structures (e.g. mushroom bodies, protocerebral bridge and central bodies) were characterized as homologous neuropile areas for chelicerate and mandibulate species. However, subsequent analysis of the spider nervous system is inconsistent with this previous view. In particular, embryonic development of Chelicerata and Mandibulata seems to differ considerably (Chabaud et al., 1990). Pross (1966) and Weygoldt (1975) describe some basic features in the embryogenesis of spiders and try to

70


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7

CRAYFISH

LOCUST abductor

\

adductor remotor

remotor TholaCiO%Xxal

promotor

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lamor axo-t-teral depressor

1

redwtor

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pmmw ant. rowor poJt rotator/

extensor flexor )M-=Pd ace.flexor bender stretcher

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FIG. 14 Structural comparison of the inhibitory MNs supplying crustacean and insect limbs. (A) Schematic representation of the leg muscles supplied by the three inhibitory neurones. Crayfish: 01, opener inhibitor; SI, stretcher-closer inhibitor; CI, common inhibitor. Locust: CII-3 common inhibitors with their targets. (B) Schematic representation of the central cells and their branching pattern. (From Wiens and Wolf (1993) .)

homologize mandibulate and arachnid brain structures. However, both authors restrict themselves to a gross morphology and fail to analyse the fate of identified neurones. Possible segmentation patterns in the supraoesophageal ganglion of spiders are discussed. But also here, no clear criteria are established which may allow the homologization of neuropile areas in the brains of mandibulates and arachnids (Weygoldt, 1975). Further, the anatomical data provided are likewise not sufficient to homologize arachnid and insect brain structures (Weltzien, 1988). Fibre


71

pathways within the arachnid brains (Breidbach, 1992; Wegerhoff and Breidbach, 1993) show an organization which deviates far from the typical structure of the neuropile of the insect brain (Mobbs, 1985). Currently, no definitive homology for the tracheate and arachnid brain structure can be suggested. Such a view is especially interesting with regard to the Onychophora brain, since some characteristics of its organization are similar to those of the spider brain (Schurmann, 1987). Thus, the onychophoran “mushroom bodies” and “central complex” (Schurmann, 1987) both seem to have their equivalence in the spider brain (Weltzien, 1988). A recent comparative analysis of an onychophoran, based on sequence analysis of 12 S ribosomal RNA, has made it highly probable, that this group is a sister group to the Chelicerata, within the arthropod phylum (Ballard et al., 1992). Future research may confirm that homologous structures exist for both the chelicerate and the onychophoran brain. If so, then this will imply that these structures represent a synapomorphy of the Chelicerata and the Onychophora. Accordingly such structures would not only be present in the brain of the common ancestor of these two groups but may even have evolved convergently to similar structures in the mandibulate brain. However, this would raise the idea that the arthropod brain had originated twice, independently! Brain structures in Chelicerata are formed according to a specific chelicerate “Bauplan”. It has been possible to establish some homologous characteristics of arachnid brain morphology allowing an assessment of a possible basic plan (Babu, 1985; Weltzien, 1988; Breidbach and Wegerhoff, 1993). Based on morphological criteria, the brain structure shows some variation for different chelicerate groups. However, this finding does not negate the idea of the existence of a common basic organization for the chelicerate brain. 3.4.1 “Primitive” Chelicerata The most primitive group of recent Chelicerata, the Xiphosura, possesses the largest brain among the Arthropoda (Patten and Redenbaugh, 1900; Patten, 1912; Holmgren, 1916; Hanstrom, 1928a; Johansson, 1933; Fahrenbach, 1977). Its main characteristics are the massive corpora pedunculata, which due to their relative position and connections appear to be homologous with the corpora pedunculata of the arachnids (Hanstrom, 1926; Fahrenbach and Chamberlain, 1987). Apart from the enormous expansion of that neuropile, the organization of the Xiphosuran brain is equivalent with that of Arachnida (Breidbach, 1992b). There are two lateral optic lobes, with a columnar-like organization (Fahrenbach, 1975). Medially, there is a sausage-formed “central body” receiving primary input from the median optic nerve (Chamberlain and

72


Wyse, 1986). Thus, the principal Xiphosuran brain areas seem to be homologous to those of Arachnida (Hanstrom, 1926). The suboesophageal ganglionic mass of Xiphosura forms a neuropile ring that encircles the oesophageus. The commissures, connecting both hemispheres, originate during early developmental stages. They form separate nerve bundles that pass ventrally around the oesophageus. The more posterior ones fuse during further development. In the abdomen a chain of five ganglia is formed. Histological (Hanstrom, 1926; Fahrenbach and Chamberlain, 1987) and immunohistochemical data (Chamberlain and Engbretson, 1982; Watson et a f . , 1984) are scarce and do not yet allow homologization with certain neurone populations in other Chelicerata. 3.4.2 Arachnida The brain of scorpions, the most primitive recent Arachnida, is formed according to the general scheme of the chelicerate brain. Hanstrom (1926) mentioned some peculiarities in the histology of the central body. But these do not subvert the principal scheme of a chelicerate brain. In an effort to characterize the functional organization of a motor system (the leg-bearing segments), the neuromeres of the suboesophageal mass of this species have attracted considerable interest (for review, see Root, 1985). However, data are not yet sufficient to homologize any of the substructures in this nerve mass with those of other Chelicerata. For the arachnids, detailed anatomical data, based on the characterization of identified neurones, are also scarce. Seyfarth et a f . (1990b) have described several serotonin immunoreactive neurones in the spider Cupiennius salei and single cell labelling studies have been performed on the same species (Brussel and Gnatzy, 1985; Eckweiler and Seyfarth, 1988; Milde and Seyfarth, 1988; Anton and Barth, 1989; Babu and Barth, 1989; Eckweiler et al., 1989; Gronenberg, 1989, 1990; Seyfarth et a f . , 1985, 1990a). Immunohistochemical studies on Cypiennius (Seyfarth et a f . , 1990b) and Opilionidae (Breidbach and Wegerhoff, 1993) show the repetition of a set of interneurones for each leg-associated neuromere. These neurones appear to be good candidates of serial homologues in true spiders and harvestmen. Comparative neuroanatomical claims for a homologous organization of brain neuropile centres in spiders (Hanstrom, l919,1921,1923,1926,1928a, 1929, 1935; Babu, 1969; Weltzien, 1988) are substantiated by comparative histochemical analysis. The distribution of acetylcholinesterase as well as the activity pattern of this enzyme in the CNS are very similar for araneid and agelenid species (Meyer and Pospiech, 1977). The distribution of acetylcholinesterase is also similar, for Salticidae and Lycosidae, but a difference in the enzyme intensity is seen for these species (Meyer and Idel, 1977). Altogether, a basically identical distribution pattern of this enzyme is apparent in different Araneae. Morphological characteristics (location,


73

pathways and termination of tracts and commissures) provide evidence of structural homologies in the neuropile organization of different Arachnida (Hanstrom, 1921, 1928a; Babu and Barth, 1984; Babu, 1985; Babu et al., 1985; Weltzien, 1988; Wegerhoff and Breidbach, 1989; Breidbach and Wegerhoff, 1993). In the suboesophageal ganglion, the principal organization of the neuropiles in Araneae (Babu and Barth, 1984; Babu, 1985; Wegerhoff and Breidbach, 1989) and Opiliones (Breidbach and Wegerhoff, 1993) is rather equivalent. Spider and harvestmen (Opiliones) are phylogenetically widely separated groups of Arachnida. The Opiliones are a subgroup of the Apulmonata which are distinct from the Megoperculata that include the Araneae (Weygoldt and Paulus, 1979a,b). Some principal morphological characteristics of Opiliones (e.g. mouthpart organization) are rather different to comparable structures in spiders. Therefore, van der Hammen (1989) places both arachnid groups in different superclasses (Rostrostomata and Myliostomata). Nevertheless, members of both these superclasses share topological and histological similarities in the organization of their nervous systems. 3.4.2.1 Suboesophageal ganglion The suboesophageal ganglion of the spider Cupiennius safei comprises 11 pairs of longitudinal tracts. There is a mid-ventral tract, a ventro-lateral tract, a centro-lateral tract, a central tract, a mid-central tract, and a mid-dorsal tract (Babu and Barth, 1984, 1989; Babu, 1985). Because of the size of their axons these tracts are interpreted as motor tracts (Babu and Barth, 1989). The other five pairs of longitudinal tracts are thought to be sensory tracts (Babu et a f . , 1985; Babu and Barth, 1989). They are located in the mid-central part of the spider’s ventral nerve mass (Babu and Barth, 1984, 1989; Wegerhoff and Breidbach, 1989). The six pairs of motor tracts of Cupiennius correspond to the 10 longitudinal tracts identified in the hemiganglion of the harvestman, Rilaena triangularis (Breidbach and Wegerhoff, 1993). Topologically, the ventral tract of Rilaena corresponds with the ventral lateral and mid-ventral tracts of Cupiennius; the lateral ventral tract of Rilaena corresponds with the central lateral tract of Cupiennius; the median tracts 2,3 of Riluenus correpond with the central tract in Cupiennius, and the lateral tracts of Rilaena correspond with the mid-dorsal tracts of Cupiennius. The architecture of the commissures is also highly equivalent in both spider and harvestman (Babu, 1985). In both arachnid groups, a dorsal, a mid-dorsal, a central, a mid-central and a ventral commissure are present (Babu and Barth, 1989; Wegerhoff and Breidbach, 1989; Breidbach and Wegerhoff, 1993). Accordingly, the principal organization of the arachnid nervous system is established by structurally conserved neuropile regions. They may be interpretated as being the homologous neuropiles of the Arachnida. Such a conclusion is substantiated by a comparative analysis of the distribution of serotonin immunoreactive neurones and CCAP-like immunoreactive neurones in

74


spiders and harvestmen (Seyfarth et al., 1990b; Breidbach, 1992a,b; Breidbach and Wegerhoff, 1993; Breidbach, unpublished). 3.4.2.2 Supraoesophageal ganglion The supraoesophageal ganglion of Arachnida is produced by the fusion of two cheliceral ganglia with the brain. The latter is characterized by a prominent input via the optic nerves which terminate in the optic mass (Hanstrom, 1926). Since in Arachnida number and position of the eyes differ, the distribution and structure of the appropriate sensory neuropiles vary for the different species (Hanstrom, 1921). Prominent neuropile regions of the spiders’ brain are: the “corpora pedunculata”, the “bridge”, “the central body”, and the “protocerebral bridge” (Hanstrom, 1926; Weltzien, 1988; Weltzien and Barth, 1991). They appear to be homologous within spiders. In a comparison of the CNS of the opilionid Rilaena triangularis with the CNS of the spider Cupiennius salei, Breidbach and Wegerhoff (1993) described the principal histological organization and identified neurones. Based on such criteria supported by structural immunological data homologous structures were claimed for both species representing different arachnid superclasses (Fig. 15). Holmgren (1916) commented on the organization of the “mushroom bodies”, which Hanstrom (1923) interpreted as being rather separate from the respective “Bauplan” in the Phalangiidae. However, when referring to the original data, presented by Holmgren, this reinterpretation appears doubtful. It becomes clear that the Gonyleptidae form a rather prominent “mushroom body” neuropile. But this does not subvert the principal morphology of their brains, which is still comparable with the scheme presented for the Phalangiid Rilaena triangularis (Wegerhoff and Breidbach, 1990). The “corpora pedunculata” of the spider Cupiennius salei (Babu, 1985) show topological characteristics (position in the ganglion, principal architecture, coupling to pathways in the brain, immunohistochemistry) similar to those of the lateral associative neuropile of Rilaena (Breidbach and Wegerhoff, 1993). Therefore, these neuropile areas should be interpreted as homologous in both species. The “bridge” of Cupiennius (Babu and Barth, 1984), likewise, corresponds with the superior rostra1 commissure in Rilaena (Breidbach and Wegerhoff, 1993). The superior associative neuropile of Riluena shows structural charcteristics (position and histological organization of the neuropile) which are comparable with the “central body” of Cupiennius. This correspondence can even be inferred at the level of identified neurones: CCAP immunohistochemistry allows the identification of four sets of individual neurones, with equal somata, pathways of their axons and projection of their arborizations. They are associated with the lateral associative neuropile in Opiliones or the “central body” of Cupiennius, respectively (Breidbach et al., unpublished). Similarly, the “protocerebral bridge” of Cupiennius can be associated with the inferior


75

A

I

‘

IBAN

I

ICCO

MAT

vFIG. 15 Diagrammatic view of the various important centres of the brains of two arachnids. (A) Spider, Cupiennius salei; view from dorsal surface showing sub- and supraoesophageal ganglia. ABDG, abdominal ganglion; 1-4 first to fourth leg ganglion; LAN, lateral associative neuropile (former “mushroom body”); OPL, optic lamella; SURCO, superior rostral commissure, former “bridge”; SUBAN, superior bilateral associative neuropile, former “central body”. (Modified from Babu (1989.) (B) Harvestman, Rilaena hiangularis; caudal view, showing the supraoesophageal ganglion. af 1,2,3, ascending fascicles 1-3; CHI, first optic chiasma; d, dorsal; IBAN, inferior bilateral associative neuropile; ICCO, inferior caudal commissure; IRCO, inferior rostral commissure; LAN, lateral associative neuropile; LAT, lateral ascending tract; MAT, median ascending tract; mf, median fascicle; NO, nervus opticus; of 1,2,3, optic fascicle 1-3; OL 1,2, first and second optic lobes; SUBAN, superior bilateral associative neuropile; SUCCO, superior caudal commissure; SURCO, superior rostral commissure. Scale bar 50 pm. (From Breidbach and Wegerhoff (1993).)

bilateral associative neuropile of Rilaena (Breidbach and Wegerhoff, 1992). Evidence for structural homologies in the brains of harvestmen and spiders is provided by a comparative analysis of pathway characteristics of serotonin immunoreactive neurones. Seyfarth et al. (1990b) described serotonin immunoreactive neurones of the nervous system of Cupiennius exhibiting a repetitive bilateral symmmetrical organization in the suboesophageal ganglia. By quantitive and qualitative criteria, this composition is comparable with that of Rilaena (Breidbach and Wegerhoff, 1993; Fig. 16). In the brain of Cupiennius, immunoreactive cell clusters were identified in locations corresponding to those described for Rilaena. Seyfarth et al.

76


FIG. 16 Serotonin immunoreactive neurones in the supraoesophageal ganglion of the harvestman, Rilaena triangularis, caudal view. d, dorsal; dm, dorsal median cluster; IBAN, inferior bilateral associative neuropile; LAN, lateral associative neuropile; mn, median neurones; OL 1,2, lst, 2nd optic lobe; pCH posterior cheliceral cluster; pdl, posterior dorsal lateral cluster; pdn, posterior dorsal neurone; SUBAN, superior bilateral associative neuropile; v, ventral; vln, ventral lateral neurones. Scale bar 100 pm. (Modified from Breidbach and Wegerhoff (1993).)

(1990b) report on a prominent laterally descending group of serotonin immunopositive neurones, terminating in the optic lobes. According to the position of the somata, axonal pathways and principal architecture, these neurones correspond to the neurones described for the harvestman Rilaena (Breidbach and Wegerhoff, 1993). Altogether, a comparative neural analysis of Araneae and Opiliones reveals an equivalent pattern not only at the level of principal neuropile architecture but also at the level of identified neurones which can be interpreted as homology. 3.5

VISUAL SYSTEMS OF ARTHROPODA

In Arthropoda the term “eye” implies not only the receptor cells per se and the associated dioptric system but also several neuropile layers, the visual ganglia or optic lobes, which may be interpreted as a lateral extension of the brain mass. The geometric arrangement of the photoreceptors (especially


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those of the compound eyes) and the central fibre system results in aesthetic structures which have attracted neuroanatomists for a long time (e.g. Kenyon, 1897; Cajal and Sanchez, 1915; Zawarzin, 1913, 1925; Hanstrom, 1924, 1928a; Exner, 1891, 1989; Strausfeld, 1976; Nassel et al., 1987). Basically, the photoreceptors of Arthropoda can be separated into two groups (see also Paulus, 1979): 1. Simple eye. This is typical of the primitive, apterygote insects, larval stages of holometabolous insects, Chelicerata, single ocelli in Myriapoda, a trio of dorsal ocelli at the frons of insects, the nauplius eye of most lower Crustacea which in several Decapoda may persist to adulthood. 2. Compound eye. Its unit is the ommatidium. Well-developed compound eyes are seen in Crustacea, Insecta and in Limulus; numerous ommatidia can be found in some Myriapoda. There are several recent accounts on the structure of the photoreceptors and associated brain structures (Wolken, 1971; Ali, 1984; Trujillo-Cen6z7 1985; Gupta, 1987a,b; Stavenga and Hardie, 1989; see also Addenda by R. C. Hardie, in Exner, 1989). Comparative studies reveal homologous structures among closely related species and groups (see also Ohly, 1975; Shaw, 1990; Strausfeld, 1990), which may even result in speculations on phylogeny (Marois and Meinertzhagen, 1990). However, a comparison of distantly related groups will show considerable differences. For example, in hemimetabolous insects photoreceptors and the associated neuropiles are generated in synchrony, while in holometabolous insects the photoreceptors of the imago develop from imaginal discs and the optic lobes stem from the larval brain. The structure of the single eyes and of the optic centres are different between Insecta and Chelicerata. Edwards and Meyer (1990) show the conservation of a specific glycoprotein associated with the arthropod compound eye; while there is a good correspondence for Insecta and Crustacea, there is some deviation when considering Lithobius (Chilopoda) . The compound eyes of Crustacea and Insecta show several distinct anatomically differences, which led Tiegs and Manton (1958) to the assumption that in both groups such a similar structure had evolved separately, a conclusion which is opposed by Paulus (1979). Owing to the wealth of information for the opposing views (see reviews or corresponding articles in bibliography, mentioned above) paired with the uncertainty of equivalence between larger phylogenetic groups, we will refrain from trying to comment on possible homologies in the visual system of different arthropod groups. 4 Conclusion

Morphological characteristics (location, pathways, termination of tracts, commissural connections) may provide evidence for structural homologies in


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the neuropil organization of different arthropod species (Yack and Fullard, 1990; Breidbach and Kutsch, 1990). Such studies are substantiated by the identification of individual homologous neurones. Homologies are the result of common cell lineages and, thus, reflect an equivalent ontogenetic programme of the appropriate cell population. Morphological criteria are sufficient to claim such a homology, when they meet the following criteria: 1. Additional to morphological criteria for such identified neurones, further information is required on biochemical and physiological specializations of such a neural set. 2. These neurones should preserve structurally conservative characteristics, both in different body segments of one species as well as in nervous systems of different species. Immunohistochemistry, which allows mapping of whole sets of identifiable serially homologous neurones, can provide information on structural and biochemical specifications of such neurones. Therefore, this technique represents a useful tool for comparative neuroanatomy, in the search for homologous neural structures. As has been pointed out above, for a rigid definition of homology such data have to be corroborated by embryological analysis. As outlined in this review, strong evidence for homology can be obtained for several neuropile centres and identified neurones.

4.1

SEGMENTATION

Segmentation of animals (metamery) is a general concept in body organization, from Annelida to Vertebrata. Apparently, this is one solution to the problem of increasing the complexity and efficiency of the whole organism (Kastner, 1965; Leise, 1990). The repetition of organs, occurring in the same number and relation, with variation of the forms, was known as a basic biological principle as early as Geoffroy de Saint-Hilaire (1807). Duplication of structures, from the genome-level to the complex morphological structures of a segment, followed by variation and modification of function, has been interpreted as “metameric logic in evolution” (Weiss, 1990). Eventually, a modular body plan is realized, which, however, may be obscured due to fusion of compartments and regional specialization during later development (French, 1990). Metamery seems to depend on the expression of homeotic genes which have been conserved throughout phylogeny (Akam et al., 1988; Pate1 et al., 1989a; Levine and Macagno, 1990; Weiss, 1990). For Arthropoda, the Antp gene family seems of great importance for segment formation. The pattern of gene deployment in early development seems to represent phylogenetically ancient roles for a specific gene and, furthermore, gene patterns appearing later in development should be explained by a recent use of this ensemble (Akam et al., 1988). The ancestral sequence of the Antp gene subfamily appears to be very similar to


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the extant Antp gene (Gehring, 1986). Therefore, it is not surprising to find a similar gene complex in the Myriapoda and Insecta. From head to tail, a different combination of these homeotic genes is used to construct the different segments (Akam et al., 1988; Weiss, 1990; French, 1990; SchmidtOtt and Technau, 1992; Sommer and Tautz, 1993). A comparison of the appropriate genes has demonstrated a far-reaching identity (homology) for Drosophila and Schistocerca (Akam et al., 1988; French, 1990). Such molecular biological studies seem to indicate that the larval mesothoracic segment may represent the ancient status of segmental organization of modern insects (Weiss, 1990). If this notion is correct, we should consider the early mesothoracic nervous system as the fundamental structure, the basic module, while all other segments have been specialized. Therefore, any search for segmental neural homologies must take into consideration the early embryogenesis with special respect to neurogenesis, and it may turn out that any structure of the mesothorax is present in any of the other segments, too, but will be suppressed early in embryogenesis or modulated throughout further ontogeny and specialization of the segments.

4.2

HOMOLOGY

The aforementioned involvement of homologous genes, apparently not only for segments of an individuum but also interspecifically, does give some hope of finding homologous structures not only as serial homologues but also among different species. This follows from the assumption that the same developmental and genetic basis will result in the same morphology (Roth, 1984; Wagner, 1989; Striedter and Northcutt, 1991). Therefore, homology should be the rule and not the exception. Based on a common “Bauplan”, epigenetic processes will modify the ontogeny which in turn is the basis of phylogenetic new lines (Miiller, 1991; Minelli and Peruffo, 1991). We have seen many examples of similarities that suggest homology at the level of reiterated segments as well as in distantly related species. Of course, similarity per se is not a proof for homology, just as dissimilarity does not exclude the possibility of homology (see section 2; Smith, 1967). Depending on individual ontogeny , sculptured by genetic and epigenetic factures, two structures (homologues) can never be 100% identical (Minelli and Peruffo, 1991). Problems may also arise when considering the possibility that specific genes may produce proteins with different functions (Baker, 1991); the expressed phenotype of neurones may vary even when they originate from the same embryonic position. On the other hand, the identification of homologous structures does not imply control by the same genes (de Beer, 1971; Striedter and Northcutt, 1991).


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4.3

PHYLOGENY

4.3.1 Independent evolution of two Arthropod brains? The present approach supports the assumption that there are two principal basic plans for arthropod brains-a cheliceral and a mandibulate one. Recent analysis of arachnid brain structure (Weltzien, 1988; Breidbach, 1991, 1992b; Breidbach and Wegerhoff, 1993) rejects previous proposals to homologize mandibulate and chelicerate brain regions. The arachnid neuropile exhibits an organization with pathway characteristics, internal morphology, immunohistochemistry, etc., which is far from the organization of the insects’ brain neuropile (Mobbs, 1985; Breidbach, 1992a). Recent comparative analysis of neurogenesis in spiders and insects outlines that the formation of the spider’s nervous system has no parallel in the development of the insect nervous system (Chabaud et al., 1990). Thus, recent embryological and anatomical data are not sufficient to homologize arachnid and insect brain structures (Weltzien, 1988). Accordingly, Breidbach and Wegerhoff (1993) propose an uncommitted terminology of the chelicerate brain indicating the considerable difference of the neural structures of Chelicerata when compared with Insecta. Such a view would imply that the Arthropoda brain does not represent a synapomorphy within this phylum. Presently, this is still a mere hypothesis, but there are some remarkable features supporting this assumption. It has been demonstrated that for Mandibulata and in Chelicerata a common groundplan of the nervous system exists. For the mandibulates, such a correponding structural organization is reflected in a reiterated expression of homeobox genes. It is not yet known whether chelicerates differ in their genetic programme of neurogenesis. A comparison with the Onychophora might elucidate some interesting aspects. As has been mentioned, molecular evidence supports the assumption that the Onychophora represent a sister group to the Chelicerata; both groups exhibit some common characteristics of their brains (compare Holmgren (1916) with Schiirmann (1987)). Accordingly, this may indicate that in terms of phylogenesis the basic arachnid brain organization is an early acquisition and possibly convergent to the groundplan of the mandibulate brain. Though neuranatomical data are still scarce, detailed embryological data (fate mapping of certain neural precursor with their progeny) and molecular data are almost non-existent. Thus, further research is needed to understand the phylogenetic relationship of Chelicerata and Arthropoda nervous systems. However, it becomes clear that the “Bauplan” within a certain group of Arthropoda is remarkably conservative, irrespective of the functional peculiarities generated by the respective neuropiles (Breidbach, 1990~).One example of such a structural conservatism has been demonstrated in the


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phylogenetic persistence of the motoneurones innervating the dorsolongitudinal muscles in Tracheata. At least in insects, a persistence of structural features of a class of motoneurones is apparent irrespective of functional changes in their peripheral targets. This notion has its parallel in the organization of identified homologous interneurones, whether for the VNC or the brain. Comparable results have been obtained for the neural architecture of the arachnid brain. In all these cases, conservation of structural features is apparent at the level of pathways and the major arborization of identified neurones. This evidence suggests that the developmental programme of the adequate nervous systems is rigid, irrespective of complex changes in function and behaviour. This is further supported by the knowledge that the neurones involved in the flight motor system of pterygote insects have their homologous partners in non-pterygote insects. It has been pointed out that early developmental programmes coincide for Insecta and Crustacea. Based on studies of the mushroom body neuroblasts (number, location, activation mode) such developmental programmes are perpetually acting even in postembryonic development. Consequently, for a comprehension of arthropod neural organization in general, these basic developmental programmes have to be deciphered. Comparative neuroanatomy provides the essentials for further progress. 4.3.2 General aspects Can our study of homology support any of the current opinions on phylogenetic relationships (see Introduction and section 2)? It is apparent that several aspects of the neural system, including the peripheral sensory system and the muscles, are very similar in all tested Insecta, from the apterygote Lepisma to the advanced fruitfly Drosophilu. Neither different developmental strategies (hemi- vs. holometabolous) nor differences in early formation of segments (short germ vs. long germ embryos; Akam el al., 1988; French, 1990; Sommer and Tautz, 1993) have resulted in an entire change of the segmental organization (with respect to the studied systems). This conservatism becomes especially apparent when comparing the CNS. In the periphery, position of sense organs, cuticle formation and even muscles, can vary. This notion is true not only for successive segments along the body axis but also when comparing different individuals, at the level of the same segment. During evolution, the CNS altogether appears to be very stable, conservative, while the periphery is more variable. Epigenetic influences are of more importance to the periphery for adapting the organism to changing requirements of the surroundings. However, more information is needed to establish the suggestion that the efferent nerves should be the most conservative insect tissue, since “as vehicles of internal communication they should be least affected by the environment” (Birket-Smith, 1984).

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W. KUTSCH A N D 0. BREIDBACH Lithobius

a

DLM

P

Lepisma-thorax Lepisma-abdomen Blattodea-thorax Saltatoria-thorax

Odonata-thorax Blattodea-a bdomen Saltatoria-abdomen Heteroptera-thorax Coleoptera-abdomen

It

Crustacea-abdomen Coleoptera-thorax Diptera-thorax Lepidoptera-thorax

FIG. 17 Schematic representation of the innervation pattern of the DLMs in different Arthropoda (excluding Chelicerata). The appropriate nerve collects axons stemming from two adjacent ganglia. Arrows indicate the possibility of transformation. c, contralateral hemiganglion; dotted line, ganglionic midline; dashed line, segmental border; dotted area, ipsilateral connective; a, anterior; p, posterior. (From Heckmann and Kutsch (unpublished).)

What would be the nearest relative of the insects to any of the other Arthropoda groups? It is obvious that the Chelicerata are far away from any of the other nervous ground plan, which would place them apart from the other groups. This coincides with all current views on phylogenetic relationship among arthropods. For Crustacea and Myriapoda correspondences can be detected with the Insecta (Fig. 17). Thereby, several systems may be structurally more equal (homologous) between Crustacea and Insecta than between Myriapoda and Insecta. Although our data are still meagre, there seems to more conformity between Crustacea and Insecta


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compared with the other pair. MNs and sensory systems have more in common for the former group-pair. This is unexpected, since systematics places Myriapoda closer to the Insecta, and both are united to form the Tracheata. An analysis of homeotic genes also places Myriapoda close to the primitive insects (Akam et al., 1988; Weiss, 1990). However, for the architecture of the centipede CNS, several dissimilarities are apparent, when compared to the insect CNS (Heckmann and Kutsch, unpublished). With respect to the centipede Lithobius, the following should be mentioned: MN somata are predominantly in the ventral position, a great part of the MN dendrites spread dorsally but also penetrate towards the ventral region; several longitudinal tracts can be differentiated; however, trials to demonstrate equivalence with the Schistocerca system have failed to date (the similarity of Crustacea and Insecta CNS structure has been exhibited by Skinner, 1985a,b). The differences between Insecta and Myriapoda with respect to the sensory system as well as neurogenesis have been mentioned above. The present comparative approach, therefore, tends to set the Myriapoda (at least with respect to knowledge, accumulated for Chilopoda) further apart from the insects than envisaged before. A recent study of the “molecular phylogeny of the animal kingdom”, based on analysis of a ribosomal RNA sequence, actually suggests a closer evolutionary distance between the brine shrimp Artemia salina (Crustacea) and the fruitfly Drosophila melanogaster (Insecta) than when Drosophila is compared with a millipede, Spirobolus marginatus (Diplopoda-Myriapoda) (Field et al., 1988).

Acknowledgements

We thank all our colleagues for sharing their interest on our comparative approach to understand the structure and function of nervous systems. We appreciate being able to read an early version of the manuscript of Drs Wiens and Wolf. Mrs A. John and Mr G. Steffens helped considerably when compiling the text. We thank Mrs R. Gimmi for taking on the burden of typing the present manuscript. The original work was substantially supported by grants from the Deutsche Forschungsgemeinschaft.

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Note added in proof

The hypothesis, that there are two principal basic plans for arthropod brains, is supported by two recent publications. Strausfeld, N. J . and Barth, F. G . (1993). Two visual systems in one brain: neuropils serving the secondary eyes of the spider CupienniG salei. J. Comp. Neurol. 328, 43-62. Strausfeld, N. J . , Weltzien, P. and Barth, F. G. (1993). Two visual systems in one brain: neuropils serving the principal eyes of the spider Cupiennius salei. J. Comp. Neurol. 328, 63-75.


prostaglandins a n d Related Eicosanoids in Insects David W. Stanley-Samuelson Department of Entomology, University of Nebraska, Lincoln, NE 68583-0816,USA

1 2 3 4 5

Introduction 116 Historical perspectives 117 Arachidonic acid metabolism: eicosanoid biosynthesis 119 Essential fatty acids in insects 127 Arachidonic and other long-chain PUFAs in insects 128 5.1 Early indications of the significance of arachidonic acid oxygenation in insects 129 5.2 The occurrence of arachidonic acid and other C20 PUFAs in insect lipids 131 5.3 Biosynthesis of arachidonic and other polyunsaturated fatty acids in insects 136 6 Physiological roles of eicosanoids in insects 147 6.1 Reproduction 147 6.2 Insect immunity 162 6.3 Fluid secretion rates in mosquito Malpighian tubules 168 6.4 Thermobiology 174 6.5 Modulation of lipid mobilization 177 6.6 Eicosanoids in invertebrate neurophysiology 178 7 Ecological significance of eicosanoids 179 7.1 Introduction 179 7.2 Blood flukes: skin penetration by cercarial larvae 179 7.3 Eicosanoids and blood-feeding 181 7.4 Prostaglandins in predator avoidance 182 7.5 Insect-derived inhibitors of eicosanoid biosynthesis 182 8 Looking ahead: desiderata and comparative eicosanoid physiology 184 8.1 Major desiderata 184 8.2 Comparative physiology of eicosanoids 197 Acknowledgements 199 References 199

Al)VAN(’ES I N INSECT PHYSIOLOGY VOI. 24 I \ H N 0-12-02422~Y

116

D. W. STANLEY-SAMUELSON

1 Introduction

Eicosanoid is an umbrella term for all of the biologically active metabolites of arachidonic and two other C20 polyunsaturated fatty acids (PUFAs). These fatty acids and their eicosanoid metabolites have manifold significance in the biology of animals (Stanley-Samuelson, 1987, 1991, 1993; StanleySamuelson and Nelson, 1993); they are generally related to two broad fields. In one, PUFAs are crucial structural components of cellular and subcellular membranes. In the other, certain PUFAs are transformed into eicosanoids, which mediate a bewildering array of biological activities, many of which centre around signalling or regulating events within cells. The biological significance of these molecules is beautifully complex. An understanding of this significance necessarily pulls us into the nutritional metabolism of lipids because most animals are not able to synthesize PUFAs de novo. Moreover, an appreciation of the roles of eicosanoids, such as prostaglandins (PGs) and related compounds, in various animal groups entails a broadly comparative physiology. Lying amid nutrition and physiology, arachidonic acid oxygenation involves three major biochemical pathways. The products of each major pathway are often themselves substrates of other pathways. The result is a plethora of oxygenated metabolites, each with a singular potential to exert a potent biological effect on cell function. The goal of this chapter is to present a historical and comprehensive review of our understanding of arachidonic acid and its oxygenated metabolites in insects and other arthropods. Research in this area dates back nearly 20 years, with quite satisfactory progress in the last 3-4 years. Nevertheless, we do not yet have a unified view of the significance of eicosanoids in arthropods, nor in other invertebrates. For this reason, after developing the major theme of the physiological roles of eicosanoids in insects in the early sections of the article, I place emphasis on models of eicosanoid action and on major gaps in our knowledge in this area. Details of the recent progress in my laboratory are covered in the last section, juxtaposed with the more obvious shortcomings. This arrangement is aimed to illuminate the rich ore that remains to be unearthed. The history and growth of this field is deeply rooted in studies of mammalian physiology, and it will be useful to develop our picture of eicosanoids in insects against the mammalian background. Insect physiology lies on the landscape of invertebrate zoology, and I will draw upon literature that extends well beyond insects to help gain a greater appreciation of the biological significance of arachidonic acid. A number of abbreviations and acronyms related to fatty acid and eicosanoid studies are in common usage. I have gathered some of these in Table 1 with the aim of providing a central and accessible lexicon.

PROSTAGLANDINS AND RELATED EICOSANOIDS IN INSECTS

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TABLE 1 A glossary of abbreviations and acronyms commonly used in eicosanoid research Term

Meaning

AA CAMP EET EFA FA GLC GC-MS HETE HPETE HPLC HX KETE LT PG PC PE PI PL PLA2 PUFA RIA TLC TRX TX

Arachidonic acid Adenosine 3’,5‘-cyclic monophosphate Epoxyeicosatrienoic acid Essential fatty acid Fatty acid Gas-liquid chromatography Coupled GC-mass spectrometry Hydroxyeicosatetraenoic acid Hydroperoxyeicosatetraenoic acid High-performance liquid chromatography Hepoxilin Ketoeicosatetraenoic acid Leukotriene Prostaglandin Phosphatidylcholine Phosphatidylethanolamine Phosphatidylinositol Phospholipid Phospholipase A2 Polyunsaturated fatty acid Radioimmunoassay Thin-layer chromatography Trioxilin Thromboxane

2 A historical perspective on essential fatty acids in vertebrates

Interest in lipid nutrition is often dated to the discovery of the fat-soluble nutrient vitamin E (Evans and Bishop, 1922). Rearing young rats on fat-free diets that were supplemented with vitamins produced a number of pathological conditions (Burr and Burr, 1929). Burr and Burr (1930) subsequently showed that the pathological conditions were ameliorated when the diets were supplemented with linoleic acid (18:2n-6). They defined those fatty acids that prevented or ameliorated the deficiency symptoms as “essential fatty acids” (EFAs; Burr and Burr, 1930). There are two groups of fatty acids that fulfil EFA requirements. Fatty acid structures and nomenclatures are presented in Fig. 1. The pathological deficiency symptoms and metabolism of EFAs in vertebrates have been considered in a number of sources (Mead, 1970; Guarnieri and Johnson, 1970; Alfin-Slater and Aftergood, 1971; Tinoco et al., 1979; Tinoco, 1982; Vance and Vance, 1985; Mead et al., 1986; Hansen, 1989-a11 referenced below as “Reviews”).

Name of Acid

Shott-Hmd Notation

Docosahexaenoic

226-3

Eicosapentaenoic

2050-3

Structure 1 3 5 7 9 11 13 15 17 19 21 % HO/ / \ / n-3

1%) into their testicular PLs. Since the C20 components were not provided in the diet, the authors surmised that a proportion of the dietary 18:3n-3 was elongated/desaturated into their C20 counterparts (Parnanen and Turunen, 1987). The 3-series PGs are produced from 20:5n-3, and the authors speculated that this may be the major PG series in this species. Eicosapentaenoic acid appears in highest proportion (9.5%) in testicular PLs of post-diapause adults (Turunen and Parnanen, 1987). Similar analyses of an artificial diet and of tissues from larval and adult Gypsy moths Lymantria dispar also point to elongation/ desaturation of dietary C18 PUFAs to C20 PUFAs (Stanley-Samuelson et al., 1992). The fatty acids of the larval medium included 18:2n-6 (about 60% total fatty acids) and 18:3n-3 (about 9Y0). The PLs of all life stages of the Gypsy moths include small proportions of 20:3n-3 and/or 20:5n-3 (about 1% of PL fatty acids). Recent analysis of PL fatty acid compositions of Malpighian tubules from the beetle Tenebrio molitor also indicates elongation/desaturation in the n-3 and n-6 families (Howard et al., 1992). Findings from the detailed research on the mosquitoes, waxmoth, cockroach and cricket species can be assembled into a broad pattern of PUFA biosynthesis in insects. Most insect species are probably represented by the waxmoth pattern, which is analogous to the general pattern that has been described for vertebrates. These species physiologically require C18 and C20 PUFAs; all requirements can be met by 18:2n-6 and/or 18:3n-3, which can be further metabolized into components that fulfil cellular needs for C20 PUFAs. Variations on this general pattern have been documented for some insect species. So far, about 15 species are known to be capable of 18:2n-6 biosynthesis by action of a novel A12 desaturase. Further elongation/desaturation of biosynthesized 18:2n-6 has been documented in two species, and in this respect, these insects are in line with the waxmoth pattern. Although a number of insects can biosynthesize n-6 PUFAs, there is no evidence that any animal can produce n-3 PUFAs. Mosquitoes are capable of fatty acid biosynthesis, but lack the desaturases that are crucial to formation of eicosanoid-precursor PUFAs. Mosquitoes are capable of retroconversion, that is, selectively shortening exogenous C20 and C22


147

PUFAs to their C18 and C20 counterparts (Dadd et al., 1987). The major implications of these remarks is that there appears to be a general pattern of PUFA metabolism among animals, and that variations on the theme have been documented in vertebrates and in insects.

6 Physiological roles of eicosanoids in insects

6.1

REPRODUCTION

The first suggestion that eicosanoids were important in the physiology of any invertebrate animal emerged from work on egg-laying behaviour in the common cricket A . domesticus, reviewed in Section 5.1. PGE2 also releases egg-laying behaviour in newly mated females of the Australian field cricket T. commodus. Work on the role of PGs in this cricket fits into the perspective of a long line of work on reproductive endocrinology and behaviour of T. commodus by Werner Loher. In this section I will recount all of our information on PG-mediated oviposition behaviour in this species, then draw upon several examples of similar work on other species of insects and other invertebrates. The emergent picture is that PGs mediate egglaying in a few insect species, certainly not most of them. PGs are probably important in other aspects of reproductive biology because they occur in the reproductive tracts of insects in which they do not mediate egg-laying. The reproductive biology of T. commodus females has been reviewed by Loher (1984). Newly emerged adults of this cricket are not sexually mature. During several days of maturation, ovaries increase in weight from about 9 mg to hundreds of milligrams. Virgin females are attracted to the male calling song, and recognition follows antenna1 contact with sex-specific pheromones. A spermatophore is transferred to females during a brief mating act. Successful mating releases profound behavioural and physiological changes in females. Two behavioural changes are reduced circadian rhythm-controlled nightly locomotor behaviour and reduced reaction to the male calling song. A third change is a great increase in egg-laying behaviour (Fig. 11). Sexually mature virgin crickets deposit a few sterile eggs on a daily basis. A successful mating releases a reiterated programme of oviposition behaviour that results in deposition of hundreds of eggs, usually in the first night after mating. After this first intense burst of oviposition behaviour, egg-laying is sustained at a level substantially higher than egg-laying in virgins. Such increases are known from many insect orders (Engelmann, 1970), however, novel features heightened interest in this system. Loher and Edson (1973) showed that the increased ovipoisition behaviour was not the result of mechanical stimulation, as known in other insect species. The effects of mating on egg production and laying in this cricket are mediated by a chemical mating-

36c

361)

320

320

280

280

240

240

‘b

.a - m %I 2 44 0 160

12 females virgin

12 females mated 200

160

#

120

80 40

1

2

3

4

5

6

7

8

9

10

days FIG. 11 The effects of mating on egg-laying by virgin and mated females of the cricket T. comrnodus. These graphs depict the numbers of eggs deposited per day. Mating (M) releases oviposition behaviour, usually during the night of copulation, resulting in deposition of large numbers of eggs.


149

factor that is transferred to females in the spermatophore. Unlike other insect species, the mating-factor originated in testes, rather than accessory glands. Extending the original work in Brady’s laboratory, Loher (1979) raised the possibility that PGE2 might release egg-laying behaviour in T. commodus. Sexually mature, virgin females were injected with 50 pg PGEz dissolved in an ethanovsaline oleo. Compared to saline-treated controls, the test females deposited about 28 times more eggs during the first day after injection (control group = 47 eggs; test group 1310 eggs). Loher (1979) remarked that further work would investigate the possibility that PGE2 is the mating factor in this species. This hypothesis was rigorously tested in a series of experiments (Loher et al., 1981). The first experiment was to quantitate PGs in spermathecae from virgin and mated females. The work showed that 100 spermathecae from mated females contained 500 pg PGE2/spermathecae, while none was detected in 100 spermathecae from virgin females. The PG was quantitated by HPLC of thep-bromophenacyl ester of PGE2 (Ganjian et al., 1981). This derivative absorbs strongly at 254 nm (extinction coefficient = 11 000/moV cm), and has a lower limit of sensitivity at about 0.5 ng. The authors did not report on the efficiency of extracting PGs from spermathecae, so the values given here are to be regarded as estimates of the amounts of PGE2 in these tissues. With respect to releasing egg-laying behaviour, the point to be made is that PGE2 was not present in spermathecae from virgins, and was present in substantial quantities in spermathecae from mated females. Similar exercises showed that PGE2 was present in spermatophores at about 20 ng/ spermatophore. This value is far lower than the 500 nghpermathecae. We concluded that PG biosynthesis takes place in spermathecae, after a factor is transferred from males to females. The next line of work addressed PG-biosynthetic activity in two tissues: spermatophore and spermathecae, The contents of these tissues were incubated with a co-factor mix (hydroquinone/glutathione/hematin) and radioactive 20:4n-6. The incubations were terminated and aliquots of extracts from the incubation mixtures were applied to TLC plates. Fractions that corresponded to PGE2 and PGFZalpha were identified by co-chromatography with authentic standards. This work showed negligible PG biosynthetic activity in spermathecae from virgin females, but substantial activity in spermatophores and in spermathecae from mated females (25-35 pmol PGE2/h/tissue). These findings suggested that PG synthesizing activity was transferred to females in the seminal fluids of spermatophores. This synthesizing also produced PGFZalpha (12-13 pmol PGFzalPha/h/tissue),which is an interesting finding, given future quantitative studies showing decreases in spermathecal PGFZalpha following mating. Arachidonic acid was not yet widely regarded as a regular component of insect lipids. This point was investigated by GLC analysis of fatty acid

150


methyl esters derived from total lipid extracts of spermathecae. Arachidonic acid comprised over 2% of total fatty acids in this tissue. These findings are the basis of the enzyme transfer model for PG biosynthesis in spermathecae of newly mated female crickets. In the work that followed these findings, this model has been thoroughly examined. The research added two inconsistencies and considerable insight into the role of PGs in releasing egg-laying behaviour in crickets. In experiments on effects of time course, numbers of spermatophore contents, arachidonic acid concentration and pH on PG biosynthesis, Tobe and Loher (1983) provided a basic characterization of the enzyme activity. The PG synthetic activity in spermatophore contents converted arachidonic acid into compounds that co-chromatographed with PGEz and PGFZalpha, always favouring formation of PGE over PGF. They showed that aspirin (Fig. 3) inhibited the enzyme activity, and estimated 50% inhibition at 55 VM aspirin. This compares favourably with the 54 p~ reported for reproductive tracts of A . domesticus (Destephano and Brady, 1977) and with values reported for dog (37 VM) and rabbit (33 WM) PG synthetase (cited in Tobe ahd Loher, 1983). PG biosynthesis was subsequently similarly characterized in the housefly M . domestica (Wakayama et al., 1986b). The inhibitor experiments reiterate an important point (Section 5.1): eicosanoid-biosynthesis inhibitors have been used to probe possible roles of eicosanoids in many studies of invertebrates. These inhibitors are functionally defined on the basis of appropriate investigations of mammalian systems, largely motivated by clinical requirements. The caveat to offer is that until the inhibitors have been characterized as eicosanoid biosynthesis inhibitors in the particular invertebrate system under consideration, they are correctly regarded as putative inhibitors (Fig. 3 ) . The pharmacological fates of these compounds in mammals and in invertebrates can be quite different. For this reason, compounds that inhibit eicosanoid biosynthesis in vitro may not be inhibitory in the whole organism. As case in point, aspirin and several other eicosanoid-biosynthesis inhibitors, do not reduce PG-mediated egg-laying behaviour in newly mated female crickets (Stanley-Samuelson and Loher, unpublished observations). There are a number of possible explanations, one of which is that the aspirin could be metabolized into an inactive form or removed from the haemolymph via Malpighian tubules before it reaches the site of PG biosynthesis, in the lumen of the spermathecae. We noted in quantitative GLC studies that spermathecae from mated crickets contained considerably more arachidonic acid than spermathecae from virgins (8 ng/organ from virgins compared to 340 ng/organ from mated females). The arachidonic acid could arise from increased elongation/ desaturation of shorter components, or it could have been imported from males along with other contents of seminal fluids. Upon analysis of spermatophore lipids by quantitative GLC, Stanley-Samuelson and Loher


151

(1983) concluded that spermatophores provided the increased spermathecal arachidonic acid. All other the arachidonic acid in spermatophores was associated with PLs. Within PL species, arachidonic acid comprised about 25% of PC fatty acids and about 4% of PE fatty acids. The high proportion of arachidonic acid in PC but not in P E suggests the arachidonic acid is accumulated into PC for special purposes (Section 5.2), in this case, perhaps for transfer to females. Once within spermathecae, the arachidonic acid could be hydrolysed from the PC and then converted into PGs. It appears that fatty acids are hydrolysed from PC and reacylated into other PL fractions within spermathecae. Unlike spermatophores, the arachidonic acid in spermathecae from mated females is about evenly distributed between PC (14%) and PE (11%). If the spermathecal arachidonic acid were not reacylated into other PL fractions, it would be expected in higher proportions in PC. Such PL metabolism depends upon spermathecal PLAz activity, which was observed in preliminary experiments (Stanley-Samuelson and Loher, unpublished observations). The sexual transfer of arachidonic acid was confirmed using radioisotope methodologies (Stanley-Samuelson et al., 1987b). Radioactive arachidonic acid (20 VCi) was injected into the haemocoels of adult males within 8 h of emergence. These males began producing spermatophores on the usual schedule for untreated individuals, 3-5 days after emergence. Spermatophores were collected and processed for total lipid extraction according to routine methods. Radioactivity in the total lipid extracts was assessed by liquid scintillation counting. Spermatophores were collected on an irregular basis for the next 49 days, all .of which contained radioactivity (mean k SE = 960 k 181 cpm, rz = 57 individual spermatophores). Radioactivity in some spermatophores was characterized on radio-HPLC that was optimized to separate fatty acid methyl esters. These analyses showed a single peak of radioactivity that corresponded with authentic arachidonic acid. Midway through this experiment, the radioactive males were individually placed into mating arenas, and allowed to mate with untreated females. The spermathecae were isolated from females at either 1 h or 24 h after mating. The amounts of radioactivity in these tissues were determined as just described. The spermathecae isolated 1 h after mating contained about 700 cpm of radioactivity, which would be expected if all the radioactivity normally found in spermatophores was transferred into the spermathecae. The spermathecae isolated 24 h after mating contained only background radioactivity; radioactivity was recovered from haemolymph and ovaries of these females. The radioactivity in haemolymph was characterized by silicic acid column chromatography. About half of it co-chromatographed with authentic PGs and half with authentic arachidonic acid. Eicosanoids are not sufficiently resolved on silicic acid chromatography to claim rigorous identification of these products. However, these results indicate that

152


radioactive arachidonic acid is transferred to spermathecae, partly metabolized to prostaglandins and subsequently moved from spermathecae into haemolymph circulation (Stanley-Samuelson et al., 1987b). Loher and Edson (1973) showed that testectomized males can produce spermatophores and mate with females, but these matings do not release egg-laying behaviour. Spermatophores from testectomized males do not contain PG biosynthetic activity, whereas spermatophores from shamoperated males contain the usual level of PG biosynthetic activity. These results indicate that formation of spermatophores with this activity depends on the testes (Tobe and Loher, 1983). Stanley-Samuelson et al. (1987b) investigated the possibility that testes are required for transfer of arachidonic acid. Males were testectomized shortly after adult emergence, then injected with 10 pCi of radioactive arachidonic acid. The treated males produced spermatophores, which were collected and assayed for radioactivity. The amounts of radioactivity in these spermatophores were similar to amounts found in spermatophores from intact males. These experiments indicate that the testes are probably the source of PG biosynthetic activity and accessory glands may be the source of the arachidonic acid in spermatophores. At the end of the 49-day experimental period the testes and accessory glands were isolated and assayed for radioactivity. The radioactivity was also characterized on radio-HPLC. Results show that these tissues accumulated and preserved very large amounts of radioactive arachidonic acid in their PLs. Since testes are not required to produce radioactive spermatophores, we speculated that accessory glands were the source. A direct test of this idea is not feasible because accessory glands are involved in producing spermatophores. The results from these experiments support the idea that PG biosynthetic activity and substrate are transferred to females by way of the spermatophores in this cricket. The increased egg-laying that accompanies mating requires that these components be transferred to spermathecae, and that PG be released into the haemolymph. When the spermathecae were removed from mated females, or the spermathecal duct was obstructed, the mating effect on egg-laying was cancelled (Ai et al., 1986). In related experiments, spermathecae from mated females were implanted into the haemocoels of sexually mature virgins. In one set of experiments, the spermathecal bulbs were purposefully injured during implantation, and in another set the bulbs were transferred intact. The virgin females that received intact bulbs continued their unmated egg-laying behaviours. The female that received the injured bulbs expressed about three-fold increases in egg laying. Data from the transplantation and spermathecae duct obstruction experiments suggest that PGs could not permeate through the spermathecal walls to the haemolymph circulation. Ai et al. (1986) put forth the idea that PGs moved along the ordinary route from spermathecae, through the spermathecal duct to the genital chamber and thence to


153

haemolymph. Sugawara (1987) controverted these results on grounds that cuticular lining in the genital chamber is an obstacle to PG movement. The sexual transfer of radioactive arachidonic acid shows that radioactivity from males can be detected in haemolymph of mated females. Granted that the exact route of P G movement remains dim, the pathway suggested by Ai et al. (1986) seems most attractive. T o gain a broader perspective on the occurrence of PGs and PG-precursor PUFAs in females of T. commodus, individual tissues from virgins and mated females were pooled, then analysed (Stanley-Samuelson et al., 1983; Ai et al., 1986). PGE2, PGF2alphaand PGA2 were quantitated in appropriate extracts by HPLC of p-bromophenacyl esters of the PGs (Table 6). These data confirmed the original quantitation of PGE2 in spermathecae from mated females. We also showed that heads, haemolymph, nerve cords and ovaries from virgins contain substantial amounts of PGE2. PGF2alpha was detected in ovaries, spermathecae and haemolymph. PGFZalphaundergoes interesting changes in titre after mating. PGFzalpha occurs at over 650 pg/ organ in spermathecae from virgin females, and, unlike PGE2, it decreases dramatically by 2 h post-mating (to under 200 pg/organ). These changes are reflected in changes in circulating titre of PGs. The ratio of PGE2/PGF2alpha changed from 0.8 in haemolymph from virgins to 2.1 in haemolymph from mated females. These changes suggest that the PG-mediated effects of mating are signalled by changing ratios of two PGs, rather than by a simple increase in haemolymph PGE2 titre. TABLE 6 Quantities of PGs in tissues from virgin and mated females of the cricket T. commodus. PGs were extracted,' then quantitated by HPLC of the p bromophenacyl esters Tissue

PGE2

Spermatheca virgin Same

n.d." n.d.

Spermatheca mated Same

500 466

Prostaglandin (per organ) PGF2 PGA2 Reference -

667

-

pg n.d.

pg pg 192 pg n.d.

Same 542 pg 232 pg Haemolymph, 100 pl virgin 10.5 pg 13.4 pg 9.4 pg Haemolymph, 100 yl mated 19.5 pg 26.0 ng 30.4 42.6 Ovary, virgin Heads, virgin

76.0 ng n.d.

n.d.

Heads, mated

130.0 ng n.d.

n.d.

a

Loher et al., 1981 Stanley-Samuelson et 1983 Loher et al., 1981 Stanley-Samuelson et 1983 Ai et al., 1986 Ai et al., 1986 Ai et al., 1986 Stanley-Samuelson et 1983 Stanley-Samuelson et 1983 Stanley-Samuelson et 1983

Not detected; dashes indicate the PG was not considered in the analysis.

al., al.,

al., al.,

al.,

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Studies of the relationship between eicosanoid structures and release of oviposition behaviour show that the effects on egg-laying are specific to Eseries PGs, and indicate an important difference from mammals (StanleySamuelson et al., 1986~).Loher (1979) originally injected 50 pg of PGE, into the haemocoels of sexually mature virgins, then observed the effects on egg-laying by counting the eggs that were deposited overnight. Nanogram doses of applied PGs also release egg-laying behaviour (Loher et al., 1981); however, these doses were applied directly to the genital chamber. We began our analysis of egg-laying by injecting five doses of PGE, into the haemocoels of virgins, then counting the eggs that were laid during the night. Figure 12 shows that although low doses cause increased egg-laying behaviour, higher doses release more egg-laying. Reasoning that exogenous PGs could be effectively neutralized by clearing them from haemolymph and by dilution within circulation, 100 pg doses were selected for all experiments. These experiments are summarized in Table 7. A few of the tested compounds (the monounsaturated fatty acid 20:ln-9, the lipoxygenase product 15-HETE, PGAl and PGI,) elicited no increase in egg-laying. PGs A, B, D and F released very little egg-laying. Higher expression of egglaying behaviour were associated with variations on PGE2 (PGEI, 6-ketoPGE1, PGE, and 15-keto-PGE,). Two other compounds (15-keto-PGFZalpha and TXB,) also elicited intense oviposition behaviour. There is no clear explanation, but higher egg-laying was generally associated with additional oxygenation of the PG backbone. The effects of the 15-keto-PGs were not TABLE 7 Relationships between eicosanoid structure and egg-laying in sexually mature virgin crickets T. comrnodus. The effects of 100 pg treatments are expressed as a percentage of the effects of mating on egg-laying behaviour. (Data derived from Stanley-Samuelson et al. (1986~)) Treatment 15-keto-PGE2 PGE2 TXB2 6-keto-PGEl PGEl PGA2 PGD2 PGFlaloha PGB; PGI2 PGAl 15-HETE

Percentage of mating effect 100 57 55 50 44 35 27 25 23 22 17 11

4 2

0

155


0

-50

I

I

I

I

I

0.01

0.1

1

10

100

pg Injected PGE, FIG. 12 The relationship between PGEz doses and egg-laying in virgin females of the cricket T. commodus. Values on the ordinate indicate increases in egg-laying relative to the low levels of egg-laying seen in untreated virgins.

156


expected on grounds that these are biologically inactive molecules in mammals. The fact that these are active in an insect bioassay indicates that the biological meaning of PG metabolism is not to be expected to necessarily follow the mammalian background. The biochemistry of PGs in insects may differ from the mammalian background on other grounds as well. In mammals, PGE2 is converted into the biologically inactive 15-keto-PGE2form by prostaglandin dehydrogenase, located mainly in lung, but also in kidney and liver. Inactivation is rapid, and most PGE, is metabolized and removed from circulation within seconds. The inactive forms can then be excreted or further metabolized (Flower, 1981). PGE2 may remain for longer periods in insect circulation. We considered this by injecting 0.02 pCi of radioactive PGE2 into the abdomen of adult virgin female crickets. After 20 min, small amounts of radioactivity were recovered from isolated tissues including ventral nerve cords, fat body and ovaries. Most radioactivity was recovered from the Malpighian tubule/hindgut complex, although substantial radioactivity remained in the haemolymph. This suggested that circulating PGs may be removed from haemolymph circulation via the normal excretory route, but at slower than mammalian rates (Stanley-Samuelson and Loher, 1985). A major shortcoming of the work is that the radioactivity was not characterized by chromatographic techniques, and it cannot be said that the injected PGE2 was or was not metabolized to another compound before excretion. There is some evidence for PG metabolism because Wakayama et al. (1986a) reported that PGs are metabolized to uncharacterized polar products by housefly tissues. This remains an important frontier of our understanding of eicosanoids in invertebrate animals. PGs may stimulate oviposition behaviour in newly mated females of some, but not all, cricket species. PGE2 releases egg-laying behaviour in sexually mature females of T. oceanicus (Stanley-Samuelson and Loher, unpublished observations). Ai and his students found that arachidonic acid and PGE2 influenced the abdominal pumping and pause duration behaviours of oviposition in the cricket Gryllus bimaculatus (Ai and Ishii, 1984; Ishii and Ai, 1985). The short-tailed cricket Anurogryllus muticus did not release egg-laying behaviour when treated with PGE2 (Stanley-Samuelson and Loher, unpublished observations). These observations fit into the context of the reproductive biology of this species, in which newly mated females do not release intensive egg-laying behaviour. PGs may release egg-laying behaviour in females of the silkmoth Bombyx mori. Yamaja Setty and Ramaiah (1980) explored this by injecting PGs into the abdomens of virgin females the day after adult emergence, then observing the number of eggs that were deposited during the following 24 h. Saline-injected controls laid 19 eggdfemale (mean, n = 12). Virgins that were treated with 1, 10 and 100 pg of PGE2 laid, respectively 32, 42 and 63 eggdindividual (mean, n > 10). These are statistically significant increases in


157

oviposition, compared to controls. There is a question on the biological significance of these numbers because normal matings result in about 500 eggdfemale. The highest PGE2 dose brought on about 12% of the number of eggs expected from mating. There is also a question of specificity because 100 yg doses of PGE, and PGFZalphaalso stimulated release of 46 and 47 eggdfemale (mean, n > 10). These results are similar to the PGE2 effects. Experiments with PG biosynthesis inhibitors (see legend, Fig. 3 ) support the idea that PGs release egg-laying behaviour in newly mated female silkmoths (Yamata Setty and Ramaiah, 1980). Males and/or females were injected with 10 pg of aspirin or indomethacin. then allowed to mate. The numbers of eggs laid during the following 24 h were then counted. Matings between untreated or control groups produced over 500 eggdindividual. Treating males and females with inhibitors resulted in substantial reduction in oviposition, by 88% with indomethacin and by 66% with aspirin. Matings between treated females and normal or control males did not reduce egglaying. Reciprocal matings, between treated males and control females, reduced oviposition to the levels observed when both sexes were treated. The authors also observed end-product reversal by injected PGE, into females that had been mated with treated males. This increased, but did not completely restore, normal egg-laying. The authors suggested that PGs, rather than PG biosynthetic activity, are transferred from male to female, resulting in increased oviposition behaviour. These data indicate that PGs are involved in releasing egg-laying behaviour. Treating virgin females with PGE, resulted in very little increase in egg-laying, while treating males with aspirin or indomethacin substantially reduced egg-laying. Aspirin and indomethacin inhibit cyclooxygenase activities in mammals, and possibly in insects. Inhibition at this point in the PG biosynthetic pathways would reduce synthesis of all PGs. It may be that PGs other than PGE and PGF are more active in releasing egg-laying behaviour. Alternatively, the quantitative relationship between PGE and PGF may important in releasing egg-laying. Their quantitative studies indicate that this may be so. The authors estimated quantities of PGE and PGF in the reproductive tracts of mated and virgin females. The PGs were extracted and partially purified on silica gel columns. Quantities of PGs were estimated using prostaglandin dehydrogenase obtained from swine lungs. Activity of this enzyme produces NADH, which the authors recorded by spectrophotometer. Their results for PGE2 indicated 15.6 pg/virgin and 29.8 yglmated female. No PGFZalphawas detected in virgins, and 13.9 yg PGF2,1ph,/female was detected in mated females. If these data represent relative amounts of these two PGs, then the ratio of PGE/PGF in female reproductive tracts changes dramatically after mating. There is convincing spectral evidence on the occurrence of PGs in the reproductive tracts of silkmoths (Yamaja Setty and Ramaiah, 1979).

158


However, it is quite doubtful that the reproductive tracts of mated females contain over 40 pg of PGs. These values are very high compared with 500 ngkpermatheca from mated females of T. commodus (Loher et al., 1981). The authors also reported 4.1 &gram tissue of PG E and 2.5 pg/gram tissue of PGF in reproductive tracts of male silkmoths, determined by the PG dehydrogenase activity technique (Yamaja Setty and Ramaiah, 1979). The authors indicated that reproductive tissues from male silkmoths express PG biosynthetic activity (Bhagya Lakshmi and Ramaiah, 1984). The biosynthetic activity may be expected. However, the quantitative values for males are also far beyond expectation, compared to 14 ng/g testes in male crickets (Destephano and Brady, 1977). Granting the high estimates in silkmoths, the reproductive tract from an individual male would have far lower amounts of PGs, certainly far too low to account for the high amounts that appear to be transferred to females. So far, the only role of eicosanoids that has been articulated in insect reproduction is the release of egg-laying behaviour in two crickets and the silkmoth. Insects have evolved many strategies of reproductive biology. A number of points argue that PG-mediated release of egg-laying behaviour is adaptive in only a few of these biological strategies. For one point, this occurs in species that do not mate until the females are physiologically prepared to deposit their eggs. For another, a number of other mechanisms lead to release of egg-laying behaviour, including mechanical stimulation and mediation by chemicals other than eicosanoids or eicosanoid biosynthetic activities. It is reasonable to conclude that PG-mediated release of egg-laying behaviour is important in an unknown, but small number of insect species. Lurking behind this remark are several reports indicating that eicosanoids are important in other, still undefined, roles in insect reproduction. Soon after the early reports on the roles PGs in insect egg-laying behaviour emerged, Murtaugh and Denlinger (1982) reported that they had detected PGE and PGFZalphain extracts from the heads and thoraces of seven insect species. The PGs were detected by commercial RIA kits, after separation by silicic acid chromatography. PG structures were not confirmed. The quantities were corrected for extraction efficiencies; however, there is no physiological context for their analyses: the insects were simply gathered from laboratory cultures before extractions. The positive points from the work are that PGs were detected in a number of common laboratory species, and that PGs were detected in testes from three species. The occurrence of PGs in reproductive tracts of insects is summarized in Table 8. I take these data to indicate that PGs occur in the reproductive tracts of most, if not all, insect species. PGs or PG-biosynthetic activity are sexually transferred to females of a number of insect species. This is so for the cabbage looper, Trichoplusia ni (Hagen and Brady, 1982), the locust Locusta migratoria (Lange, 1984), the


159

TABLE 8 PGs occur in the reproductive tracts of these species Species A . domesticus

Mechanism of transferring PGs to females Reference

T. commodus Locusta rnigratoria Bombyx mori

Enzyme Enzyme Enzyme Transfer

transfer transfer transfer PGs

Trichoplusiu ni Musca domestica Sarcophaga crassipalpis T. molitor

Mechanism not clear Substrate transfer Not tested Not tested

Destephano & Brady, 1977 Loher et al.. 1981 Lange, 1984 Yamaja Setty & Ramaiah. 1984 Hagan & Brady. 198.2 Wakayama et 01.. 1985 Murtaugh & Denlinger. 1982 Murtaugh & Denlinger. 1982

housefly Musca domestica (Wakayama etal., 1986a, b) and the haematophagus bug Triatoma infestans (Brenner and Bernasconi, 1980). Increased oviposition behaviour does not necessarily follow the PG transfers. I will review the details of this work, then place these observations into the broader context of the roles of PGs in reproductive biology. The central point that PGs may be involved in many aspects of reproduction will be underscored by drawing upon literature from other invertebrate animals. Compared to virgins, the reproductive tissues of mated females of the cabbage looper Trichoplusia ni contain increased quantities of PGE2 and PGF2,,lpha(Hagen and Brady, 1982). The PGs were determined by RIA of extracts from all reproductive tissues except ovaries, corrected for extraction efficiency. Based on the results from another lepidopteran, the silkmoth (Section 2.1.2), it might be thought that PGs, rather than PG-biosynthetic activity, are sexually transferred in T. ni. Studies on PG biosynthetic activity were not reported, but it seems that the PG increases are due to transfer of PG biosynthetic activity. First, the reproductive tracts from virgin males and females had about the same amount of PGEl (about 25 pghissue equivalent); tissues from mated females contained almost three times more (71 pgkissue). Second, compared to virgins, tissues from mated females contained twice as much PGFlalpha(16 vs. 32 pghissue equivalent), but none was detected in reproductive tissues from virgin males. Although other mechanisms are possible, these increases are parsimoniously explained by transfer of PG biosynthetic enzyme during mating. If this is so. then PGs are transferred differently in silkmoths and the cabbage looper moth. Treating virgin females with 100 pg injections of P G E , , PGEl and PGF22,1ph:, did not increase egg-laying. PG-biosynthetic activity is sexually transferred to females of Locusta migratoria (Lange, 1984). Using the assay of Loher et al. (1981). Lange showed that PG-biosynthetic activity was present in spermatophores. opalescent glands and seminal vesicles of adult males; activity was not

160


detected in accessory glands other than the opalescent glands, nor in testes. Low levels of activity were detected in spermathecae of virgins, which increased by an order of magnitude after mating. PG-precursor PUFAs were not considered. Again, PG treatments did not stimulate egg-laying in sexually mature virgin locusts (Lange, 1984). Male and female houseflies feature low proportions of arachidonic acid in PLs (Wakayama et al., 1985). Houseflies also sexually transfer eicosanoids, albeit by transferring precursor PUFAs, rather than biosynthetic enzymes or PGs per se. In experiments by Wakayama et a f . (1986a), female houseflies were allowed to mate with males that had been prelabelled with radioactive arachidonic acid. The amounts of radioactivity recovered from females 4 h after mating were consistently 3-5 times more than the amounts of radioactivity in male reproductive tracts. The authors suggested that males mobilize internal arachidonic acid during their lengthy mating periods. The fatty acid is transferred to male reproductive tracts, and subsequently transferred to females. Most of the transferred radioactivity was incorporated into lipid fractions within females, and small proportions were converted into several PGs, including PGE2, PGF2nlphaand PGA2/B2. The possibility that these PGs may release egg-laying behaviour remains unexplored. Mating-stimulated increases in PGs have been observed in the reproductive tracts of female insects, as just described for representatives of Orthoptera, Lepidoptera and Diptera. Such increases may occur in many, or even most, insect species. So far, it appears that eicosanoids may not increase in females of the German cockroach Blattella germanica (Casas et al., 1986). Virgin females of this species have substantial quantities of PGs, and they do not appear to increase upon mating. These analyses were carried out on whole animals, which may obscure small changes in PG titres within female reproductive tracts. As we have just seen, one biological effect of these increased PG titres is the release of egg-laying behaviour in some species. The following examples from lower vertebrates and invertebrates will help develop a broader view of the significance of PGs and other eicosanoids in invertebrate reproduction. The point to be taken is that it can be reasonably speculated that PGs play important, and far ranging, roles in the reproductive biology of all animals. Viewed in this context, the significance of eicosanoids in insect reproduction can be expected to go far beyond the effects of PGs on egg-laying behaviour in a limited number of insect species. The original discovery of PGs comes from work on the reproductive biology of mammals. As a point of history, PGs take their name from the prostrate gland, and they are found in seminal fluids of all mammals that have been appropriately analysed. These molecules are of tremendous biological and clinical importance in mammalian reproduction. PGs are also active in reproductive biology of fishes and reptiles.


161

Ovulated female goldfish release F-type PGs into the water. These PGs constitute the goldfish postovulatory pheromone and they stimulate male spawning behaviour (Sorensen et al., 1988). Other arachidonic acid metabolites appear to mediate release of gonadotropin in goldfish (Chang et al., 1989). Similarly, arginine vasotocin induces PG biosynthesis in reproductive tracts of the viviparous lizard Scefoporus jarrovi (Guillette et af., 1990). The PGs are thought to contribute to stimulation of oviductal contractions associated with parturition in reptiles. PGFZalpha titres approximately doubled in the blood of female garter snakes by about 15 min following mating (Whittier and Crews, 1989). The increased PG titres are thought to have a functional role in inducing post-mating declines in sexual behaviour. These examples are not meant to form an exhaustive review. They are lifted from the literature to illustrate the point that eicosanoids are involved in diverse aspects of vertebrate reproduction. Eicosanoids modulate reproductive physiology and behaviour of invertebrates beyond Insecta. The hatching factor of the barnacle Bafanus balanoides was identified as the novel eicosanoid 10,11,12-trihydroxy5,8,14,17-eicosatetraenoicacid (Holland et al., 1985). Similarly, 8-HETE is the hatching factor of another barnacle, Eliminius modestus (Hill et al.. 1988). Aside from Arthropoda, eicosanoids figure in many aspects of molluscan reproduction. Among bivalves, Osada et al. (1989) implicated PGs in spawning in the scallop Patinopecten yessoensis. PGs are also involved in gastropod reproduction. PGE2 increased egg production in the snail Helisoma duryi (Kunigelis and Saleuddin, 1986), and PGs accelerate the onset of egg laying in the pond snail Lymnaea stagnafis (Clare et al., 1986). Among Echinodermata, the eicosanoid (8R)-HETE, induces oocyte maturation in some, but not all, starfish (Meijer et a f . , 1986). In their detailed studies of the sea urchin model of fertilization, Schuel el a f . (1985) indicated that eicosanoid biosynthesis is one of the redundant mechanisms involved in preventing polyspermic fertilizations. A prominent (1 1R) and (12R) lipoxygenase activity occurs in eggs of the sea urchin (Hawkins and Brash, 1987). These examples indicate that eicosanoids are major regulators of reproductive biology in vertebrate and invertebrate animals. These regulators act at organismal, tissue, cellular and subcellular levels. Expression of some eicosanoid functions, such as release of egg-laying behaviour and ovulation pheromone signalling, is likely to be restricted to a relatively small number of animal species. Other roles, such as prevention of polyspermic fertilizations, appear to be fundamental actions that may occur throughout Animalia. When viewed on the broader landscape of animal reproduction. it is not surprising to see that eicosanoids occur in the reproductive tissues of insects. Exploring the biological meaning of these compounds will undoubtedly reveal hitherto unseen levels of regulation in the physiology of insect reproduction.

D.W. STANLEY-SAMUELSON

162

6.2

INSECT IMMUNITY

Eicosanoids play central roles in the inflammatory and immune responses of mammals (reviewed in volumes edited by Goodwin, 1985 and Levine, 1988). Among other important roles in immunity, eicosanoids potentiate inflammatory reactions such as vasodilation, which induces oedema, and biosynthesis of complement proteins. Many eicosanoids, including cyclooxygenase and lipoxygenase products, mediate hypersensitivity reactions in mammals. Eicosanoids are involved in natural killer cell anticancer actions. Besides their roles as mediators of immune responses, PGs act as physiological modulators of immune functions in mammals. The importance of eicosanoids in mammalian immunity responses is underscored by recognitions of their growing importance in clinical settings. Beyond mammals, the roles of eicosanoids in immunity of nonmammalian vertebrates are coming to light (Rowley, 1991). Given the importance of these molecules in the immunity of most vertebrate classes, it may be that mediation of immune responses is another area of animal biology in which eicosanoids are fundamentally and intricately involved. Based on such reasoning, Stanley-Samuelson et al. (1991) put forth the view that eicosanoids are mediators of haemocytic immunity in insects and, by extension, other invertebrates. To place this work in a broader perspective, the volumes edited by Gupta (1986, 1991) and Brehelin (1986) and the recent reviews by Ratcliffe (1985), Dunn (1986), Boman and Hultmark (1987), Lackie (1988) and Karp (1990) provide the following sketch of invertebrate immunology. Invertebrate immunity is an amalgam of several distinct physiological systems that provide protection from potential pathogens. Bacterial infections induce insects to synthesize a number of antibacterial proteins. These include lysozymes, enzymes that directly attack bacteria by hydrolysing the peptidoglycan cell walls of bacteria. Insects also produce a number of antibacterial proteins that function by disrupting bacterial cell membranes. These include cecropins (4 kDa), attacins (21-23 kDa), diptericins (8 kDa) and insect defensins (4 kDa). One of the proteins induced in the silkmoth Hyalophora cecropia was designated P4. This protein is one of the first proteins that bind to bacteria. Recently Sun et al. (1990) recognized that this protein belongs to the immunoglobulin superfamily of proteins. They renamed the protein haemolin, and suggested that it takes part in formation of a protein complex that initiates the immune response. Insects also produce agglutinating proteins, lectins, which may serve to agglutinate invading micro-organisms. Antibacterial proteins are regarded as part of the immune response to bacterial infections. Recently, however, Russell and Dunn (1991) showed that lysozyme is released into the pupal midgut lumen of M . sexta in the absence of bacterial infection. The authors suggested that the high midgut


163

lysozyme activity confers prophylactic protection from bacterial infections during metamorphosis. In addition to the humoral immunity conferred by various antibacterial proteins, insects elaborate several forms of cellular, or haemocytic. immunity. Gupta (1991) regards granulocytes and plasmatocytes as immunocompetent cells, which he calls “immunocytes”. Other haemocytes are not considered to be immunocompetent. This is only the latest of many interpretations of insect haemocytes. Given the understanding that haemocytes from different phylogenic groups are not uniform in morphology and that the biological roles of separate categories, and subcategories, of haemocytes are not clearly delineated, consensus on the nomenclature of insect haemocytes is unlikely in the near future. Fortunately, the broader issues of cellular immunity are well understood. When they are challenged with foreign materials, haemocytes respond in any of three ways: phagocytosis, encapsulation and nodule formation. Beside these direct interactions with invaders, haemocytes indirectly contribute to immune responses. These include haemolymph coagulation at wound sites, release of the prophenoloxidase activating system and detoxification reactions. Haemocytic actions are the immediate responses to bacterial infections. Antibacterial proteins are not detectable in haemolymph until 6-12 h after infection. For example, when living bacteria were injected into larvae of the tobacco hornworm, M. sexta, the bacteria were rapidly removed from the circulating haemolymph. The clearance was accomplished through cellular actions. Nodule formation was the prominent cellular action during the first 2 h (Dunn and Drake, 1983), followed by phagocytosis during the next 6 h (Horohov and Dunn, 1983). These differences in rcsponse time courses can be used experimentally to dissect humoral and haemocytic responses to bacterial infections. Figure 3 indicates that each of the major eicosanoid biosynthetic pathways may be probed by selective pharmaceutical inhibitors. These inhibitors were used to test the idea that eicosanoids are involved in mediation of cellular actions that clear bacterial infections from the haemolymph circulation of tobacco hornworms M . sexta (Stanley-Samuelson et al., 1991). I emphasize, again, that use of such inhibitors is based on the assumption that they exert similar actions in mammals and invertebrates. Although we argued that this was probably so for the inhibitors we employed, the pharmacology of such compounds in any invertebrate has not been elucidated. In preliminary experiments, test larvae were treated with 50 pg of dexamethasone ( PLA2 inhibitor) and control larvae were treated with ethanol. Then both groups were infected by intrahaemocoelic injections of a red-pigmented strain of the insect pathogen Serratia marcescens (c. 5 x lo5 bacterial cells/larva). Haemolymph samples were withdrawn, diluted and streaked on agar plates at 15, 30, 45 and 60 min post-infection. Figure 13 shows that no detectable bacteria were recovered from the control larvae, while large numbers of bacteria were recovered from the test insects.


164

15

30

45

60

Time (Minutes) FIG. 13 Recovery of the bacterial pathogen Serratia marcexens from tobacco hornworm haemolymph samples. Values on the ordinate represent the number of colony-forming units (cfu) of bacteria in haemolymph samples. Test larvae (+) were first injected with dexamethasone, and control larvae 0 were first injected with ethanol. Both groups of larvae were then intrahaemocoelically infected with c. 5 x 10' bacterial cells. A t 15 min intervals after infection, 15 pl haemolymph samples were serially withdrawn from the caudal horn, diluted, and plated on agar plates. (Data from Stanley-Samuelson et al. (1991).)

The 50 pg dexamethasone treatments that were injected in these initial experiments are high, and potentially misleading dosages. The effects of far lower doses, from 1.4 X lop5 to 0.14 pg/larva, were subsequently examined. The experiments were similar to those just described, with the exception that haemolymph was sampled once, 60 min post-infection. Figure 14 shows that increased dexamethasone dosages were associated with increased bacterial recovery. Inhibition of eicosanoid biosynthesis appeared to compromise the ability of larvae to clear bacteria from their circulating haemolymph. It may, on these grounds, be expected that similar inhibition of eicosanoid biosynthesis would also result in increased larval mortality. This idea was tested by treating groups of test larvae with the dexamethasone dosages just


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Dexamethasone Concentration (ugharva) FIG. 14 Dose-response relationship between dexamethasone treatments and bacterial recovery. M . sexta larvae were first injected with the indicated doses of dexamethasone, and then infected as described in Fig. 13. Haemolymph samples were withdrawn and plated as described in Fig. 13 after 1 h. Statistically significant differences in recovery of bacteria are indicated by letters above each column. (Data from Stanley-Samuelson et al. (1991).)

described, then injecting them with bacteria. Figure 15 shows that increased larvae mortality attended the increased dexamethasone dosages. These data indicate that immunocompromised larvae, by inhibition of eicosanoid biosynthesis, experience higher mortality due to bacterial infections. Eicosanoid biosynthesis depends upon availability of free substrate, which for the most part is hydrolysed from membrane PLs by action of PLA,. Dexamethasone is thought to indirectly inhibit eicosanoid biosynthesis by exerting its effects upon PLA2 activity. Figures 13-15 indicate that dexamethasone compromises the cellular immune responses to bacterial infections. Figure 15 also shows that the dexamethasone effect was reversed by treating the larvae with arachidonic acid. In these experiments, larvae were treated with the highest dose of dexamethasone (0.14 pg/larva), then treated with 50.0 pg of arachidonic acid. These larvae were then infected with bacteria. The arachidonic acid treatments significantly reduced larvae mortality. These end-product reversal experiments add considerable support to the idea that eicosanoids are crucial mediators of insect cellular immunity. Dexamethasone inhibits all eicosanoid biosynthesis through its action on PLA2 activity. The effects of dexamethasone do not allow us to draw


166

60

50

A

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40

30

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a

20 10

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Concentration Dexamethasone (@larva)

FIG. 15 Dose-response relationship between dexamethasone and larval mortality. Tobacco hornworms were injected with the indicated doses of dexamethasone or dexamethasone plus arachidonic acid (ARAC). and then infected as described in Fig. 13. After 14 h, surviving larvae were counted. Statistically significant differences in survivorship are indicated by letters above each column. (Data from StanleySamuelson et ul. (1991).)

conclusions on the relative importance of individual eicosanoid biosynthetic pathways (Fig. 3) in cellular immunity. We probed the cyclooxygenase and lipoxygenase pathways by treating separate groups of test insects with indomethacin (cyclooxygenase inhibitor) and esculetin (lipoxygenase inhibitor). The results of these experiments showed that inhibition of each pathway impaired the hornworms' ability to clear bacterial infections from their haemolymph; however, the the degree of impairment did not match the dexamethasone results (Fig. 16). We interpreted these results to suggest that products of both major pathways were involved in the organismal response to bacterial infections. Indeed, when larvae were treated with a combination of indomethacin and esculetin, the results were identical to the outcomes of experiments with dexamethasone (Stanley-Samuelson et af., 1991). We return, again, to the point that the occurrence and biological significance of arachidonic acid in insects is not yet widely appreciated. When eicosanoid mediation of an important physiological action is educed,


167

1

n=15 T

0.9

n=14

T

0.8

n=15

T T

n=16

0.7 0.6 0.5 0.4

0.3

0.2

n=24

n=12

T

T

0.1

0

ETOH control

ESC +

DEX

ESC

INDO

MAL

INDO

.

Treatment

FIG. 16 Effects of eicosanoid biosynthesis inhibitors on tobacco hornworm immune to bacterial infections. Test larvae were first injected with dexamethasone (DEX), indomethacin (INDO), esculetin (ESC), maleic acid (MAL) or a 1:l combination of esculetin and indomethacin. Control larvae were first injected with absolute ethanol (EtOH). Larvae were then infected as described in Fig. 13. After 2.5-h incubation. haemolymph samples were withdrawn and plated. The number of individual larvae per treatment is given above each column, the height of which represents the proportion of larvae from which any bacteria were recovered; error bars represent 1 SE of the mean. (Data from Stanley-Samuelson ef al. (1991).)

it is necessary to address basic issues of eicosanoid biosynthesis and eicosanoid-precursor fatty acids. Preliminary evidence on these points was presented (Stanley-Samuelson et al., 1991). Ogg et al. (1991) studied the fatty acid compositions of lipids prepared from the haemocytes of M . sexta larvae. These studies indicated that the fatty acid compositions of haemocytes are different from the serum lipids and from the insects’ culture medium. Small quantities of eicosanoid-precursor fatty acids, 20:3n-6, 20:4n6 and 20:5n-3, were detected in total lipids, phospholipids and phospholipid fractions. Although the amounts were quite low, they were consonant with

168


the emergent picture of fatty acid compositions of terrestrial insects (references in Section 5.2). The haemocytes were also able to incorporate exogenous radioactive arachidonic acid into cellular phospholipids (Ogg et al., 1991). The radioactive arachidonic acid was selectively incorporated into individual phospholipid species. Most radioactivity was incorporated into PC, less into PE and still less into PS/PI (these two fractions co-chromatograph on onedimensional TLC). These differences in uptake are at least partly due to the differential amounts of individual phospholipid species within cells (Bridges, 1983). In lepidopterans so studied, the choline and ethanolamine fractions are the predominant phospholipids, and phosphatidylcholine occurs in higher proportions than does phosphatidylethanolamine. Similar differences in incorporation of eicosanoid-precursor polyunsaturated fatty acids have been recorded in mammalian cell systems (references in Ogg et al., 1991), and these differences are taken to indicate that the proportions of eicosanoid precursors in phospholipid species are individually regulated within cells. The idea that eicosanoids mediate the cellular immune responses of an invertebrate animal opens a novel and very broad field for investigations. Some of these investigations are on-going projects in my laboratory; recent results will be presented in the context of major gaps in our understanding of eicosanoid systems (see Section 8.1).

6.3

FLUID SECRETION RATES IN MOSQUITO M AL P I G H I A N TUBULES

Modulation of the events that control ion and water transport may be one of the fundamental roles of eicosanoids in metazoan animals. The mammalian kidney is the best understood system in this respect (Bonvalet er al., 1987). Products of the cyclooxygenase, lipoxygenase and epoxidase pathways play a number of important physiological roles in kidneys, many of which are not fully understood. For examples, PGE2 decreases sodium and chloride reabsorptive rates in the medullary thick ascending limb and collecting tubule (Bonvalet et al., 1987). The lipoxygenase product 12-HETE reduces glomerular filtration rates in isolated kidneys (Quilley and McGiff, 1990). Escalante et al. (1991) indicated that epoxygenase products also act in kidney physiology by affecting the sodium-potassium-2 chloride transporter. These examples are presented to make the point that eicosanoids may be expected to be involved in a number of organ- and cellular-level actions in transport physiology. Beyond mammals, eicosanoids also are involved in ion and water transport in amphibians. The short-circuit current across the skin of the frog Rana temporaria is increased by prostaglandins. This is due to increases in active sodium transport in epithelial cells (Barry et al., 1975). PGE? stimulated chloride, sodium and potassium secretion in the skin glands of


169

another frog, Rana esculenta (Bjerregaard and Niellsen, 1987). PGs increase hydrogen ion excretion, but not ammonium ion transport, in the urinary bladder of the toad Bufo rnarinus (Frazier and Yorio, 1990; Yorio et al., 1991). Although we have a far weaker grip on the roles of eicosanoids in transport physiology in invertebrates, these compounds are likely to be of great significance. Freas and Grollman (1980) showed that hypoosmotic stress stimulated PG biosynthesis in gill tissues of the marine bivalve Modiolus dernissus. PGs also regulate sodium transport in the freshwater bivalve Ligurnia subrostrata. PGE2 inhibited sodium influx, and caused no change in sodium efflux. Inhibitors of PG biosynthesis increased sodium influx (Saintsing and Dietz, 1983). The early indications that eicosanoids are involved in transport physiology of insects was discussed earlier (Section 5.1). Taken together, the information from mammals, other vertebrates and invertebrates support the idea that regulation of transport physiology may be a fundamental eicosanoid action in the Metazoa. The Malpighian tubules and rectum comprise the renal organs of the majority of insect species. The longstanding wisdom holds that a primary urine is formed by secretory processes in the Malpighian tubules. The primary urine is considerably modified before excretion by resorptive events in the rectum. These fluid secretion and resorption processes are under neuroendocrine control (Spring, 1990). Diuretic hormones and related factors increase fluid secretion in Malpighian tubules and decrease resorption of water in the rectum. Antidiuretic hormones and such factors act on the rectum to increase fluid .resorption. Antidiuretic hormones are generally thought not to act on the Malpighian tubules, although this view is called into question by the recent suggestion that antidiuretic hormone also acts on the Malpighian tubules of the cricket A . domesticus (Spring et a f . , 1988). Petzel and Stanley-Samuelson (1992) put forth the hypothesis that eicosanoids regulate aspects of Malpighian tubule physiology in insects. The idea was examined in a series of experiments that are logically parallel to the experiments just described for the work on insect immunity. We reasoned that if eicosanoids were involved in regulating basal fluid secretion rates, then inhibiting eicosanoid biosynthesis would result in changes in fluid secretion. We examined fluid secretion rates in Malpighian tubules from adult females of the yellow fever mosquito Aedes aegypti using routine Ramsay assays (Petzel et al., 1987). The results of these experiments are summarized in Fig. 17. The arachidonic acid analogue eicosatetraynoic acid (ETYA) completely blocks eicosanoid biosynthesis because it inhibits PLA2 as well as cyclooxygenase and lipoxygenase. ETYA significantly reduced fluid secretion rates (Fig. 17). Although these data suggest that eicosanoids are involved in Malpighian tubule physiology, as discussed earlier, they provide


170

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FIG. 17 A summary of the effects of 100 +M eicosanoid biosynthesis inhibitors on fluid secretion rates in in vitro preparations of Malpighian tubules from the mosquito A . aegypti. EtOH, ethanol; ETYA, eicosatetraynoic acid (PLA2 inhibitor); ECS, esculetin (lipoxygenase inhibitor); 525A, SKF 525A (cytochrome P-450 inhibitor); indo, indomethacin (cyclooxygenase inhibitor); nap, naproxin (cyclooxygenase inhibitor).

no insight into the possible significance of specific eicosanoid biosynthetic pathways (Fig. 3). We probed the lipoxygenase pathways with esculetin, which had no effect on basal fluid secretion rates. The epoxygenase pathways in mammals are inhibited by SKF-525A (Escalante et al., 1991). This compound produced a small, statistically significant, decrease in fluid secretion, indicating that epoxygenase products may be active in regulating an unidentified aspect of the process. These products are certainly active in kidney and vascular systems in mammals (McGiff, 1991). Inhibition of the cyclooxygenase pathways with indomethacin produced results similar to the effects of ETYA (Fig. 17). These findings indicate that PGs are important agents in regulating fluid secretion physiology.

171

PROSTAGLANDINS AND RELATED ElCOSANOlDS IN INSECTS

Three points add support to the idea that PGs are involved in regulation of fluid secretion. First, indomethacin expressed its effects in a dosedependent manner: over the range of 0.1, 1.0, 10.0 and 100.0 pg treatments, fluid secretion rates decreased with increasing indomethacin doses (Fig. 18). Second, naproxin, another cyclooxygenase inhibitor of dissimilar chemical structure, produced results similar to the indomethacin effects. Third, in all experiments, indomethacin did not inhibit CAMP-stimulated fluid secretion. Taken together, these findings strongly support the hypothesis that eicosanoids, especially PGs, are important elements in fluid secretion physiology. The roles of eicosanoids in mosquito Malpighian tubule physiology are linked to Dadd’s discovery that arachidonic acid, or certain other structurally related polyunsaturates, are essential for normal development of mosquitoes. In his original research on mosquito nutrition, Dadd assessed

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rhdomethacin] FIG. 18 Dose-response relationship between the indicated doses of indomethacin and fluid secretion rates. Each bar represents the mean percentage change in fluid secretion rate (SE). (Data from Petzel and Stanley-Samuelson (1992).)

172


normal growth in terms of a flight index based on the ability of adult mosquitoes to support themselves in free flight (Dadd, 1983a). The flight indices were taken to represent unidentified, albeit crucial, physiological failures that were expressed as a weakened flight capability. Two points indicate that one biological role of the dietary arachidonic acid is related to eicosanoid biosynthesis. First, addition of pharmaceutical eicosanoid biosynthesis inhibitors to the defined larval growth medium mimicked the effects of arachidonic acid deficiency on flight indices (Dadd and Kleinjan, 1984). The effects of the inhibitors appeared to be expressed through their actions on arachidonic acid metabolism because increased levels of dietary arachidonic acid attenuated the inhibitor effects. Second, supplementation with PGFZalpha,but not PGE2, partially compensated for essential arachidonic acid deficiency (Dadd and Kleinjan, 1988). Dadd and Kleinjan concluded that dietary arachidonic acid is required by mosquitoes for multiple physiological functions, one of which is providing substrate for prostaglandinogenesis. Our identification of a role for PGs in Malpighian tubules (Petzel and Stanley-Samuelson, 1992) can be taken as just one physiological significance of essential arachidonic acid in mosquitoes. The occurrence of eicosanoids in the Malpighain tubules of other insect species would add support to the idea that these molecules arc involved in Malpighian tubules generally. Stanley-Samuelson and Loher (1986) detected PGEZ in Malpighian tubules of the cricket T. commodus. These findings arc to be regarded as tentative because the PGs were detected by RIA of unfractionated extracts of whole Malpighian tubules. The structures of the PGs have not been confirmed by GC-MS analysis. However, Howard and Stanley-Samuelson (1990) confirmed the presence of arachidonic acid, and other eicosanoid precursors, in Malpighian tubules from the beetle T. molitor. We recently employed immunohistochemical methods to investigate the presence of PGE2 and PGF2alphain selected insect tissues, including Malpighian tubules (Witters, 1991; Howard et al., 1992; Petzel et al., 1993). This methodology has the advantages of working with single tissues taken from individual insects of known physiological status, and the ability to localize antigens within organs. There arc also major pitfalls associated with immunohistochemical localizations. Potential problems can be minimized by developing a series of appropriate controls (Beltz and Burd, 1989). Plate I shows immunohistochemical staining of PGE2 in Malpighian tubules from the mosquito A . aegypti. 6.3.1 Integrating endocrine and eicosanoid regulation of cellular events Given that transport functions in insects are regulated by diuretic and antidiuretic hormones and related factors (Spring, 1990), the regulatory

PLATE 1. Immunohistochemical visualization of PGE? in Malpighian tubules from the mosquito A . aegypti (photograph by N . A . Witters). After incubations with primary and secondary antibodies, the PG was localized by staining with horseradish peroxidase. The repeating staining pattern suggests that principal, but not stellate. cells contain PGEz.



173

actions of eicosanoids may not be intuitively clear. How are the actions of eicosanoids integrated with other known regulators of cell function? Cellular biosynthesis of eicosanoids may represent an additional tier of regulatory events. Bonvalet et al. (1987) suggested three types of interactions between eicosanoids and hormone regulators of renal function. In the first place, eicosanoids directly influence ion transport. This is quite likely in mosquito Malpighian tubules. Results of in vitro studies suggest that the Malpighian tubules in adult female mosquitoes can exist in three states, namely basal state, inhibited state and stimulated state. The tubules maintain a low, basal level of fluid secretion activity. The Malpighian tubules can be stimulated to higher rates of fluid secretion by diuretic factors (Petzel et al., 1987); basal fluid secretion is inhibited by other factors (Hayes el af., 1989). PG biosynthesis inhibitors reduced basal fluid secretion rates, indicating that prostaglandins directly influence transport activities in the basal secretory state. At a second level of integration, several hormones alter PG biosynthesis in mammalian kidneys, some stimulating and other inhibiting biosynthesis. We have no data on the effects of diuretic and antidiuretic hormones on eicosanoid biosynthesis in insects. Information on this point is clearly a major desideratum. Third, PGs modulate the effects of hormones, in particular by attenuating the hydroosmotic effect of antidiuretic hormone in mammals. These modulating actions occur through receptor-mediated changes in intracellular cAMP levels, and they imply interactions among hormone signal transduction pathways. Petzel (1993) and Stanley-Samuelson and Petzel (1993) propose that three main signal transduction pathways are active in Malpighian tubules (Fig. 19). These are: (1) the adenylate cyclase pathway, which directly increases intracellular CAMP; (2) the PLA2 pathway, which leads to release of arachidonic acid and subsequent biosynthesis of eicosanoids; and ( 3 ) the phospholipase C pathway which produces two intracellular signals. One is inositol-3-phosphate which leads to increases in intracellular calcium and the other is diacylglycerol which up-regulates protein kinase C. Our current model (Fig. 19) suggests that products of each of these pathways can impact upon the other pathways, and that the final cellular response is the outcome of the interactions of these pathways. We have preliminary data on interaction between the adenylate cyclase and PLA2 pathways: PGE2 stimulates increases in intracellular cAMP (Parish, Petzel and Stanley-Samuelson, unpublished). A detailed report of these findings is forthcoming.

174


arachidonic acid

IP3

DAG

phospholipid phospholipase A2 FIG. 19 Potential interactions among three main transmembrane signal transduction pathways.

6.4

THERMOBIOLOGY

6.4.1 Mediation of behavioural fevers Many mammals express fevers as one line of defensive response to infections. Fever also occurs in non-mammalian vertebrates, although the increased body temperatures may be mediated by behavioural, rather than physiological, mechanisms. Behavioural fever has been observed in frogs (Casterlin and Reynolds, 1977; Myhre et al., 1977), a lizard (Bernheim and Kluger, 1976) and several fishes (Reynolds et al., 1976). There are no data on the role of eicosanoids in febrile responses in insects. Here I present information from other invertebrates which raises the speculation that eicosanoids mediate fevers in insects. Some invertebrate animals, including insects, express behavioural fevers. Following bacterial infection, the freshwater crayfish Cambarus bartoni produced a 2°C fever by moving into a zone of warmer water (Casterlin and Reynolds, 1978). This response may be mediated by endogenous PG biosynthesis because increasing doses of PGE, induced similar fevers in uninfected individuals. Three marine arthropods, the American lobster Homarus americanus, the


175

pink shrimp Penaeus duorarum, and the horseshoe crab Limulus polyphemus, similarly increased their body temperatures after injections of 100 pg of P G E l . Two terrestrial arthropods, the scorpions Bathus occitanus and Androctonnus australis, produced remarkable fevers after treatment with physiological doses of PGEl (Cabanac and Le Guelte, 1980). These arthropods regulated their body temperatures by placing themselves in selected positions along temperature gradients in a sand box. PGEl treatments resulted in 15-20°C increases in temperature preferrenda. The data in these cases are not conclusive on the roles of PGs in mediating fevers in invertebrates. For the most part, the idea is supported by observations of increased body temperatures after treatments with individual doses of P G E I . In a previous review of this area (StanleySamuelson, 1987), I raised a number of questions, answers to which might help place our appreciation of this work on firmer ground. Do the appropriate PGs naturally occur in these species? Is the fever response specific to PGEI? Does PG biosynthesis increase after infection? These points remain open, but they may assume considerable interest if induction of febrile responses is a major component of invertebrate immunity. 6.4.2 Body temperature set points in desert cicadas Many insect species utilize behavioural and physiological means to maintain constant body temperatures in parts of their bodies while they are exposed to a wide range of ambient temperatures. Many animals, including a large number of insect species, regulate their body temperatures by behavioural means, such as basking in the sun (Casey, 1981). Queen bumblebees provide particularly fetching examples of physiologically regulating the temperatures of parts of their bodies (Heinrich, 1979). When flying at higher temperatures, the queens regulate the patterns of haemolymph circulation to favour transfer of heat from the thorax, where it is produced by the prodigious flight muscle energy metabolism, to the abdomen, from whence it can be radiated away from the body. During flight at low temperatures, haemolymph circulation is regulated such that heat is not transferred out of the thorax. Beyond regulating thoracic temperatures during flight activity. during the early days of the colony cycle, queens enhance developmental rates by incubating their brood clump. Heat is generated in the flight muscles, which are uncoupled from wing movements. Again, haemolymph circulation is regulated to favour movement of heat from the thorax to the abdomen. The abdomens are adpressed to the brood clump during incubation, which facilitates transfer of a high proportion of the heat to the developing brood. Albeit with some differences in specific pattern, many of the insects that physiologically regulate their thoracic temperatures do so by altering haemolymph circulation. Some insects reduce their body temperatures by evaporative cooling.

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Cicadas are unique in their ability to regulate their body temperatures by transporting water across their cuticles, which cools the animal by evaporation from the body surface. This sweating has been analysed in some detail in cicadas of genera Diceroprocta and Tibicen (Toolson, 1987; Toolson and Hadley, 1987; Hastings, 1989; Hastings and Toolson, 1991). These cicadas allow their body temperatures to rise to a particular level, at which point they actively transport water via specialized tissues to their body surfaces. The temperatures at which water transport begins are called the temperature set points. Toolson, Ashby, Howard and Stanley-Samuelson (unpublished) suggest that eicosanoids play a number of roles in the physiology of sweating in cicadas. The cicadas were treated with inhibitors of PG biosynthesis by intrahaemocoelic injection, then observing the effects of the compounds on the temperature set points and water loss rates. Treating cicadas with the cyclooxygenase inhibitors aspirin and paeonol resulted in significantly higher water loss rates and lower thoracic temperatures, compared to control animals. Treatment with l-series PGs produced a small, but significant, increase in temperature set points. Injecting the precursor to l-series PGs, 20:3n-6, increased water loss rates, but did not impact body temperature. These results indicate that PGs are mediators of thermoregulatory functions in normothermic cicadas. Body temperature is the outcome of a complex network of events. In cicadas, body temperature is thought to minimally involve sensing and integrating body temperature information, control of thermoregulatory behaviours, regulation of circulation patterns, and control of transport events within sweat glands. Future work is aimed at understanding possible roles of eicosanoids within these systems. Because eicosanoids may be involved in a number of these areas, a direct relationship between eicosanoid biosynthesis and organismal-level temperature regulation is not expected. As in many other insect species, the occurrence and metabolism of eicosanoid-precursor fatty acids has only recently been considered in cicadas. GC-MS analysis confirmed that the PLs of selected tissues, including thoracic muscles, abdominal dorsal glands, Malpighian tubules, guts and ovaries include small proportions of 20:3n-6, 20:4n-6 and 20:5n-3 (Stanley-Samuelson et al., 1990a). These small proportions are consistent with findings from other terrestrial insects (Section 5.2). Male and female adult cicadas were able to convert exogenous radioactive arachidonic acid into three PGs, PGD2, PGE2 and PGFZalpha, as well as other unidentified products (Stanley-Samuelson et al., 1990a). The radioactive products were identified by a single TLC step, so the PGs should be regarded as tentatively identified. None the less, we have sufficient evidence to postulate that eicosanoids are important in the thermobiology of these cicadas.


6.5

177

MODULATION OF LIPID MOBILIZATION

We return, for a moment, to our discussion of the interactions among eicosanoids and other regulators of cell function (Section 6.3.1). Several hormones, including glucagon and epinephrine, stimulate mobilization of lipids in mammalian adipose tissue. As described for kidney function, the effects of the hormones are modulated by locally produced PGs: PGE2 reduces the amount of lipid that is mobilized in response to epinephrine (Mead et al., 1986). The PG effect is thought to be exerted through action on adenylate cyclase. Two studies indicate that PGs similarly modulate lipid mobilization in insects. A number of reviews detail the roles of adipokinetic hormone in maintaining homeostasis of haemolymph lipids (van der Horst, 1983; Beenakkers et al., 1985; Kanost et al., 1990). One adipokinetic hormone action is the release of diacylglycerol from fat body cells. In preliminary studies with in vitro locust fat body preparations, Wagemans, van der Horst and Stanley-Samuelson (unpublished) examined the interaction of PGE2 with adipokinetic hormone. Fat body preparations were treated with 10 pmol of adipokinetic hormone and increasing doses of PGE2, then the amount of lipid released into the medium was assayed. In these experiments, addition of adipokinetic hormone resulted in the release of 1004 pg of lipid. When added to the fat body preparations in doses from 5x to 5 x lop3, PGE2 attenuated the hormone effects, as shown in Table 9. These results support the hypothesis that PGs modulate lipid mobilization in insects. Yamaja Setty and Ramaiah (1982) presented results indicating that PGEl decreased lipid mobilization in male pupae of the silkworm B. mori. In their experiments, male pupae were treated with saline, PG E l or with PGE isolated from B. mori male reproductive organs. Three parameters were assayed: esterase activity, lipase activity and haemolymph lipid titre. TABLE 9 Effects of five doses of PGE2 on amount of lipid released in response to a standard treatment of adipokinetic hormone. Experiments were performed on in vitro preparations of L. rnigratoria fat bodies. (Unpublished preliminary experiments by Wagemans, van der Horst & Stanley-Samuelson.) PGE2 treatment

Lipid released

(M)

(I%)

x 10-5 x x x lo-' x 10-3

1004 415 314 245 190 94

0 5.0 2.5 5.0 2.5 5.0

178


Compared with controls, pupae that were treated with PGE, expressed significantly reduced fat body esterase and lipase activities, and they had significantly lower levels of haemolymph lipid. These experiments were not designed to examine the possible relationships between adipokinetic hormone and PGs, but they do suggest the attractive idea that PGs are one of the elements that regulate lipid metabolism in insects. These reports suggest that PGs may regulate lipid metabolism in two ways. One is the direct down-regulation of lipid mobilization as just described, and the other is through interaction with adipokinetic hormone. As in other potentially important physiological roles of eicosanoids in invertebrates, a number of questions remain open. Do insect fat bodies produce PGs? Work on another line (Howard et al., 1986; Jurenka et al., 1986; see Section 7.5) indicates that PGs and possibly other eicosanoids are biosynthesized by microsomes prepared from fat bodies of the cockroach P. americana. Given that fat body is generally competent to biosynthesize eicosanoids, and the PGEl impacts on lipid metabolism, further investigation of the roles of eicosanoids can be expected to yield important insights into regulation of lipid metabolism in insects.

6.6

E I C O SA N O I D SIN INVERTEBRATE N E U R O P H Y S I O L O G Y

PGs have been detected in extracts of brains from crickets (StanleySamuelson et al., 1983) and from ticks (Stanley-Samuelson and Lane, unpublished), as well as in extracts of heads and thoraces from seven other common laboratory insect species (Murtaugh and Denlinger, 1982). StanleySamuelson and Loher (1986) (Section 6.1) put forth the idea that in releasing egg-laying behaviour in the Australian field cricket, PGE-, exhibits a hormonal mode of action. We suggested that egg-laying behaviour results from the interaction of PG with still unidentified elements of the cricket nervous system, although the nature of the interaction is not clear. Nanda and Ghosal (1978) observed that treating females of the cockroach P. arnericana with exogenous PGF7alphnresulted in reduced amounts of neurosecretory material in the neurosecretory cells of the pars intercerebralis. These workers injected 1.25 pg PGF-,;,l,,,Janimal, the injection site unspecified, then prepared the brains of the cockroaches for lightmicroscope observations. They suggested that PGs are involved in the physiology of secretion in these cells. Aside from these ideas, no direct PG roles in insect neurobiology have been suggested. However, findings from the sea hare Aplysia indicate that eicosanoids are fundamental elements in synaptic physiology. The peptide FMRFamide (a tetrapeptide denoted by single letter amino acid code) is active in sensory neurons of Aplysia. This molecule produces effects opposite to the effects of 5-HT, that is, it hyperpolarizes the nerve


179

cell membrane, decreases action potential duration, and increases the outward potassium current, among other actions. Piomelli et al. (1987a,b) showed that arachidonic acid and the lipoxygenase metabolite 12-HPETE (Fig. 6) mimic the actions of FMRF. They demonstrated that neural tissue biosynthesizes lipoxygenase products, and that inhibition of phospholipase activity suppresses the FMRF effects. The authors concluded that the release and metabolism of arachidonic acid mediate the inhibitory response to FMRF in Aplysia sensory neurons. Further characterization of the 12lipoxygenase pathway showed that the 12-HPETE is metabolized to 12-keto5,8,10,14-eicosatetraenoicacid (12-KETE). The authors suggested that 12HPETE and its metabolite 12-KETE participate in synaptic actions in the Aplysia nervous system (Piomelli et al., 1988). To the extent that Aplysia is taken to model the details of neurophysiology in animals, these results suggest that eicosanoids are fundamentally involved in synaptic transmission. From the point of view of our efforts to gain a thorough appreciation of insect physiology, studies of this molluscan model argue that, as discussed elsewhere, eicosanoids appear to represent a hitherto unseen tier of regulatory elements.

7

7.1

Ecological significance of eicosanoids INTRODUCTION

Many of the ecological relationships between and among populations of organisms are mediated by chemical means. In this section I want to describe briefly three ecological relationships that are mediated by eicosanoids, then review a short series of papers on eicosanoid biosynthesis inhibitors reported in insect defensive secretions. I will then try to make the point that insects may be able to impact on the physiology of other organisms by elaboration of eicosanoids and compounds that inhibit eicosanoid biosynthesis. These findings may be of considerable interest in understanding the relationships between insects and other organisms.

7.2

BLOOD FLIJKES: S K I N PENETRATION B Y C E K C A K I A L .L A R V A E

I reviewed the broader roles of eicosanoids in blood flukes elsewhere. from which these abbreviated remarks are taken (Stanley-Samuelson, 1087. 1991). Eggs of the blood fluke Schistosoma mansoni (Fig. 20; Platyhelminthes, class Trematoda) leave their mammalian hosts along with the faeces. If they are deposited in water, the eggs hatch and continue larval development in snails. After leaving their intermediate hosts, free-swimming cercarial larvae infect mammalian hosts by ingestion with drinking water or by burrowing through the skin.

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FIG. 20 A cercarial larva of the blood fluke Schistosoma mansoni. Eicosanoids mediate many aspects of the host-parasite relationship between this parasite and its mammalian hosts. In behavioural biology, eicosanoids educe a cessation of swimming and penetration of host skin. Eicosanoids regulate larval transformation to adult forms. Adult blood flukes biosynthesis eicosanoids, which may reduce efficacy of host defence mechanisms. (I thank B. Salafsky for the gift of this photograph.)

Among skin lipids, free PUFAs are most efficacious in stimulating cercarial penetration (Austin et al., 1972). Cercarial penetration is the outcome of two behaviours, cessation of swimming and initiation of penetration. Both behaviours are mediated by eicosanoids, rather than by the PUFAs themselves. In studies of cercarial penetration into membranes prepared from EFA-deficient and EFA-replete rats, there was about three time less penetration into membranes from the EFA deficient rats (Salafsky et al., 1984). Another line of experiments used eicosanoid biosynthesis inhibitors. Interperitoneal injections of the cyclooxygenase inhibitor ibuprofin led to time-dependent accumulation of the drug in the skin of EFA-replete rats. Cercarial penetration of membranes prepared from these rats was reduced. The proportion of inhibition was correlated to the amounts of ibuprofen that accumulated in the skin (Salafsky and Fusco, 1985). Under conditions that reduce the levels of PUFAs and reduce eicosanoid biosynthesis, cercarial penetration is reduced.


181

Cercariae are competent to synthesize eicosanoids. When cercariae were incubated with radioactive linoleic acid, radioactivity was recovered in HPLC fractions that corresponded with PGE2, PGD2, LTC4, LTB4 and 5HETE. Fusco et al. (1986) concluded that formation of eicosanoids is an essential step in penetration of human skin by cercariae of S. mansoni. In this instance, their action in a host-parasite relationship is expressed in the behavioural physiology of the blood fluke. In addition to their roles in skin penetration, eicosanoids are formed and secreted by adult blood flukes within mammalian hosts (Salafsky and Fusco, 1987a,b). The authors speculated that the worms utilize the immunomodulatory actions of eicosanoids as a mechanisms to evade mammalian host defence systems. Based on the thorough discussion of PGs as immunoregulators in mammals by Plescia and Racis (1988), I regard this as a quite reasonable suggestion. Eicosanoids appear to be involved in three aspects of the relationship between blood flukes and their mammalian hosts, specifically cessation of swimming, initiation of penetration behaviour and the possible modulation of the host immune system by worm-derived prostaglandins.

7.3

EICOSANOIDS AND BLOOD-FEEDING IN TICKS

PGs have been detected in the saliva of cattle ticks, Boophilus microplus. by bioassay and chromatography techniques. Higgs er a f . (1976) reported PGlike activity by assaying extracts of tick saliva on the rat stomach strip, the chick rectum and rat colon. The biologically active material corresponded to PGE2 on TLC. Dickinson et al. (1976) reported that the muscle-contracting substance was inactivated by incubation with PG-dehydrogenase. There are other reports of PGs in tick tissues aside from the salivary glands; however, the physiological roles of these compounds remain unclear. Kemp and Bourne (1980) considered the possibility that PGs might alter the tick behaviours associated with attachment to their hosts. They concluded that histamine may result in detaching from the hosts, but other mediators, including bradykinin, PGE2 and 5-HT, had no effects on the behaviours. Ribeiro et al. (1988) determined that the saliva from the tick Zxodes damimini contained substantial amounts of 6-keto-PGFlalph,, the stable degradation product of prostacyclin. The appearance of 6-keto PGFlalphais accepted as evidence for the occurrence of prostacyclin. The authors suggested that prostacyclin may help tick feeding in several ways. First, by preventing platelet aggregation, prostacyclin could prevent host haemostasis, and thus maintain blood flow from injured vessels to the tick’s mouthparts. Second, by preventing mast cell degranulation, prostacyclin could minimize the oedema that attends tick rejection reactions. Third, the prostacyclin could increase the amount of blood available for tick feeding by


182

inducing hyperaemia. These points should be taken as quite reasonable hypotheses that remain to be tested. If they are supported by future endeavours, it may well turn out that eicosanoids are important players in the relationship between haematophagous arthropods, including insects, and their hosts.

7.4

PROSTAGLANDINS IN PREDATOR AVOIDANCE

The first discovery of an eicosanoid in any invertebrate animal came from work on the gorgonian octocoral Plexaura homomella (Weinheimer and Spraggins, 1969). This, and a very few other, coral species contains extraordinarily high amounts of PGA2 and its ester derivatives, as much as 8% of the wet tissue weight. Most octocoral species do not feature unusual levels of eicosanoids. Gerhart (1991) summarized his view that the high levels of PGs play a major role in octocorals, not necessarily in physiology, but as chemical defence agents against predation by coral fish. His argument is based on two main points: first, many predators develop a learned aversion to emetic foods, and second, orally delivered PGs induce vomiting in wide range of vertebrates, including a number of fish species. Gerhart observed that octocorals which contain high levels of PG or food pellets that were laced with PGs induce vomiting in five species of fish, and that the fish form a learned aversion to the coral and to the treated food pellets. The aversions to the emesis-producing foods are formed after a single episode of vomiting. These data suggest that the high levels of PGs found in certain octocorals may serve to protect the coral from predation. This idea is attractive, and if it is supported by further behavioural experiments that relate the amount of PG in the food to aversions, it will stand as an example of eicosanoidmediated interactions between populations.

7.5

INSECT-DERIVED INHIBITORS OF EICOSANOID BIOSYNTHESIS

A number of natural plant products inhibit biosynthesis of various eicosanoids. A now-classic example is the cyclooxygenase inhibitor aspirin. This is a natural product from members of the Salicaceae. Insects put forth a large number of chemical defences in their ecological interactions with other populations of predators, parasites and microbial pathogens (Blum, 1981). Most of these chemicals are toxic or noxious compounds, not generally thought to interact with regulators of physiological processes. Recently, Howard et al. (1986) raised the idea that certain of the chemicals found in insect defensive secretions may exert their effects by inhibiting PG biosynthesis.

183


Two aromatic ketones, 2'-hydroxy-4'-methoxyacetophenone and 2'hydroxy-4'-methoxypropiophenone, from the defensive secretions of the red flour beetle Tribolium castaneum, are potent inhibitors of cyclooxygenase preparations from mammals and insect tissues (Howard et af., 1986). Their results show that the insect-derived compounds are about 15 times more active than aspirin when assayed against cyclooxygenase from mammals. They are about equal in activity against the insect cyclooxygenase. This difference in sensitivity marks an important point, because it indicates that there are substantial differences in the enzymes from mammals and insects. A number of other insect defensive components are also cyclooxygenase inhibitors, summarized in Table 10 (Jurenka et at., 1986, 1989). Insectderived cyclooxygenase inhibitors have been found in secretions from Hymenoptera, Hemiptera and Coleoptera. What is the biological significance of these compounds? Inhibition of cyclooxygenase activity may be taken as an adventitious effect of chemicals that serve other, currently undefined, roles. Alternatively, these compounds may mediate aspects of chemical ecology through their effects on eicosanoid biosynthesis in predators or other worrisome species. We have just seen that eicosanoids mediate the host-parasite relationship between schistosome cercariae and their mammalian hosts. It is not unreasonable to suppose that species equipped with a packet of eicosanoid biosynthesis inhibitors are less useful to parasites or predators. Ribeiro and Sarkis (1982) presented a model of how an insect-derived inhibitor of eicosanoid action may function in a host-parasite relationship. These authors showed that the saliva of the blood-sucking bug Rhodnius prolixus contains a thromboxane antagonist. This antagonist is one of a TABLE 10 Insect-derived PG biosynthesis inhibitors. (Compiled from Howard er al. (1986), Jurenka et al. (1986, 1989)) Compound

2'-Hydroxy-4'-methoxypropiophenone 2'-Hydroxy-4'-methoxyacetophone Methyl anthranilate o-Aminoacetophenone Methyl salicylate 2,s-Dihydrophenylacetic acid-lactone Salicylaldehyde 2,6-Di hydroxyacetophenone 2,4,6-Trihydroxyacetophenone 5,7-Dihydroxy-2-nonylchrome 1-(2,6-Dihydroxyphenyl)dodecan-l-one 2,4-Dihydroxyacetophenone In bovine cyciooxygenase inhibition assay

Comparison to equal dosage of aspirin"

> Aspirin > Aspirin > Aspirin =

Aspirin

> Aspirin > Aspirin =

Aspirin

< Aspirin =

Aspirin

< Aspirin > Aspirin > Aspirin


184

number of compounds in the saliva that serve to help R . prolixus acquire and process a blood meal. TXA2 (Fig. 5) mediates platelet aggregation, which is an essential step in blood coagulation. Two lines of evidence are convincing on the point that Rhodnius saliva certainly interferes with haemostasis (Ribeiro and Garcia, 1981). First, bugs without their salivary glands are far less effective in taking a blood meal. Second, Rhodnius saliva markedly increased bleeding time when applied to small wounds in rat tails. To the extent that the thromboxane antagonist component of the salivary secretion reduces haemostasis in the host, the ability to modulate eicosanoid actions is an important aspect of this host-parasite relationship. The role of PGs in egg-laying in certain insects was detailed in Section 6.1. Eicosanoids are involved in many aspects of mammalian reproduction, and I suggested that eicosanoids may similarly be involved in insect reproduction in ways that remain to be identified. A ramification of this idea is that inhibition of eicosanoid biosynthesis may reduce fecundity in some insect species. This is also supported by studies with the insect growth regulation buprofezin. Uchida et al. (1987) showed that this compound, a chiten synthetase inhibitor in larval insects, reduces fecundity in adults of the brown rice planthopper Nilaparvata lugens. The authors showed that buprofezin reduced PG synthesis and reduced egg-laying in adults in a dosedependent manner. Injecting PGE2 into buprofezen-treated females reversed the buprofezen effects on egg-laying. The effects on this compound on fecundity appear to be through its effects on PG biosynthesis in N . lugens. Again, the chemical ecology of eicosanoid biosynthesis inhibitors may include the influence of eicosanoids on fecundity.

8 Looking ahead: desiderata and comparative eicosanoid physiology

8.1

MAJOR DESIDERATA

8.1.1 Discovering new eicosanoid actions in insect physiology The main thrust of this chapter is that eicosanoids bear in an important way on many areas of insect biology. In its current form, our knowledge base is uneven and thinly spread. Plainly, a first hope is that we gain new knowledge of eicosanoid actions in insects. By this I mean a broadening of information. If PGs regulate aspects of ion and fluid transport in mosquito Malpighian tubules, what about the Malpighian tubules of other insects? What about other epithelial transport phenomena, such as the processes in recta? What about other secretory processes? Similar questions attend each of the findings reviewed in this chapter. Another aspect of broadening our information base relates to the many areas of insect physiology, hitherto unconsidered in this connection, in


185

which eicosanoids may figure prominantly. Just as an example to facilitate thought on the topic, Robertson (1986) considered the roles of eicosanoids in regulating insulin secretion from pancreatic beta cells. Cyclooxygenase and lipoxygenase products modulate insulin secretion in opposite ways. Although many questions remain, it is clear that an understanding of pancreatic beta cell function in healthy and in diabetic individuals will include appreciation of the roles of eicosanoids. This example from mammalian physiology illustrates several key points. First, endocrine physiology can be modulated by the roles of eicosanoids in releasing or secreting hormones. Second, eicosanoids can modulate the physiological effects of other eicosanoids. Third, the uptake and metabolism of circulating energy metabolites can be regulated, in part, by locally formed eicosanoids. These points relate to insect physiology in this way: the endocrine regulation of organismal events in insects may be similarly compounded by eicosanoid systems. In this connection, many areas of what is now viewed as classical insect physiology may be more thoroughly understood by considering the potential actions of eicosanoids in any given physiological event.

8.1.2 Mechanisms of eicosanoid actions A second desideratum is pegged to understanding how eicosanoids exert their actions. The discussions of Malpighian tubule physiology (Section 6.3) and interaction with adipokinetic hormone (Section 6.5) lead to the idea that eicosanoids function through their interactions with specific receptors. Halushka et al. (1989) reviewed current knowledge of eicosanoid receptors in mammalian cells. They point out that although receptor studies represent a very important element in the picture of eicosanoid action, they lag far behind the progress in other areas of eicosanoid biochemistry and physiology. This is also true in invertebrate studies. Freas and Grollman (1981) described specific binding of PGA2 in gill tissues of the marine bivalve, M . demisus, which remains the only published work on the topic in any invertebrate. Their experiments were based on classic competitive binding procedures. Radioligand binding studies require considerable amounts of tissue, which may present a substantial barrier to progress in such tissues as mosquito Malpighian tubules. I concur with Halushka et af. (1989) that this is an area in which tools of modern molecular biology would be especially useful. Smith (1989) presented a general biochemical mechanism of eicosanoid action, as understood in mammalian studies. In his view, all eicosanoid actions are expressed through G-protein-linked eicosanoid receptors (Fig. 21). This idea is unifying, and helps elucidate the interactions between eicosanoids and other regulatory molecules (Section 6.3, 6.5). Smith produced a two-part model of eicosanoid action. First, eicosanoids are local hormones that function to co-ordinate effects of other hormones. Second,

186


Eicosanoid,

(PG,Tx,LT)

Cellular Response FIG. 21 A general model of biochemical mechanisms of eicosanoid actions. Eicosanoid interact with receptors that are specific to individual structures. The receptors are coupled to G proteins, which cause changes in intracellular levels of second messengers and ions through their interactions with effector proteins. (Derived from Smith (1989).)

eicosanoids function through G-protein-linked receptors. This view implies that different G proteins are influenced by individual ligand-occupied receptors. The G proteins variously stimulate and inhibit adenylate cyclase, activate phospholipase C, open or close calcium channels and impact fluxes of other ions. The biological effects of eicosanoids and other regulatory molecules may be integrated into a single cellular expression if there are several types of receptors, each coupled to a different G protein. Continued efforts in this area will greatly magnify our understanding of eicosanoids in insect physiology.

8.1.3 Advances in the biochemistry of eicosanoids in insects Our appreciation of the significance of eicosanoids in insect biology is limited by important lacunae in our knowledge of eicosanoid biochemistry. Use of pharmaceutical compounds that inhibit eicosanoid biosynthesis yields insights into possible eicosanoid actions. Selective inhibition of cyclooxygenase, epoxygenase and lipoxygenase pathways (Fig. 3) may help


187

determine which major group of eicosanoids are active in a particular setting. Experiments of this sort do not produce information on which of many eicosanoids mediate a specific action. Progress in this more detailed enquiry will depend on careful analysis of eicosanoid biosynthesis in specific tissues. O n a broader theme, we still do not know how well insect eicosanoid systems will fit the background from clinically driven research on mammals. Does eicosanoid biosynthesis depend on PLA2 action? Are eicosanoid actions mediated by specific receptor proteins? Are all the eicosanoids known from mammals produced in insects? D o insects and mammals exhibit similar eicosanoid catabolic pathways? We are investigating two of these points in our laboratory. In the following paragraphs I present some of our recent findings, with special emphasis on current shortfalls. 8.1.3.1 Phospholipase A2 activity According to mammalian orthodoxy, PLA2 is the rate-limiting enzyme in eicosanoid biosynthesis (Dennis, 1987). Insect lipase and phospholipase activities have received very little attention. Lipase activities were recorded in a few studies of insect tissues, including larval waxmoths Galleria mellonella, cockroaches P. americana, and the locust L. migratoria (Wlodawer and Baranska, 1965; Male and Storey, 1981; Hirayama and Chino, 1987). These and similar studies were concered with energy metabolism, emphasizing hydrolysis of neutral, rather than polar, lipids. Phospholipases were detected in a number of dipterans (Bieber et al., 1969; Hanumantha and Subrahmanyam, 1969; Isharaza et al., 1970; Kumar et al., 1970), however, we have virtually no information beyond determining the presence of these activities. More to our point, these findings were reported long before the significance of eicosanoids was unveiled, and we have no insight on hydrolysis of arachidonic acid. We are investigating PLA2 activity in homogenates prepared from fat body of tobacco hornworms (Uscian and Stanley-Samuelson, 1993). This work is based on a standard assay of PLA2 activity (Reynolds et al., 1991). Phosphatidylcholine with radioactive arachidonic acid esterified to the sn-2 position is prepared in the form of vesicles. Fat body homogenates are prepared by two centrifuge steps (1310 x g for 10 min, then 11 750 x g for 2 min). The supernatant protein is incubated with radioactive substrate. Total lipid extracts are separated into PLs and free fatty acids on TLC. The amount of radioactivity in each fraction is determined by liquid scintillation counting. The fat body preparations contain a PLA2 activity that can hydrolyse 20:4n-6 from PLs. The activity is sensitive to protein concentration, time, temperature, and pH (Fig. 22). PLA2 activity was not present in cell-free serum, and was inactivated by boiling the preparation for 10 min. An unusual feature of the fat body PLA2 activity is that it is calciumindependent or else it requires very low calcium concentrations. PLA2 in each class has been described from mammals. Almost all of the PLA2

188


activities and purified enzymes require millimolar calcium concentrations for optimal activity. Calcium-independent PLA,s are known from some mammalian sources, including secretory granular membranes from rat parotid gland, guinea pig intestinal brush border membrane and rat jejunal brush border membrane (Pind and Kuksis, 1987; Gassama-Diagne et al., 1989; Mizumo et al., 1991). A newly recognized class of PLA2s, known as high molecular weight PLA2, has been described from a number of mammalian sources (van den Bosch et al., 1990). These are soluble enzymes that require calcium concentrations in the range of 10-7-10-6 M for activity, far below the millimolar ranges required by most of the described PLA2s. The information that we have gained so far convincingly shows that a PLA, activity that can hydrolyse 20:4n-6 from PLs is present in the fat body. Several points remain to be addressed. First, PLA2 is not the only pathway to release the sn-2 fatty acid from PLs. One alternative pathway is the sequential action of PLAl and lysophospholipase. These steps sequentially remove the sn-1 and the sn-2 fatty acids. Another pathway begins with phospholipase C, which hydrolyses the phosphate base moeity, leaving diacylglycerol. The fatty acids can be released from the glycerol backbone by lipases. Given these alternatives, it is appropriate to refer to PLA, activity rather than PLA2. Purification of the PLA, activity will help settle this issue. On the second point, the PLAz activity we detected is not clearly linked to eicosanoid biosynthesis. One point that would help explore such linkage is the substrate specificity of the enzyme. Detection of an arachidonyl-specific PLA, would provide an indication that the PLA2 is important in making eicosanoid-precursor PUFAs available. There is so far only one other report on PLAz activity in conjunction with eicosanoid biosynthesis in insects. Reproductive tissues from the firebrat Thermobia domestica contain a PLA2 activity that can hydrolyse PUFAs from the sn-2 position of PLs (Ragab et al., 1992). This work was couched in terms of two previous papers that report lipoxygenase activity in these tissues (Ragab et al., 1987, 1991). The authors indicated that the activity they described was linked to eicosanoid biosynthesis because the PLA? inhibitor ETYA reduced eicosanoid biosynthesis. Howcver, this point is confounded because ETYA also inhibits eicosanoid biosynthesizing enzymes. We also note that the rate of the PLA, activity in the firebrat tissue homogenates is remarkably high, by three to four orders of magnitude, compared to the rates found in similar preparations of mammalian tissues (Gronich et al., 1988; Schalkwijk et al., 1989) and compared to our preparations from tobacco hornworm fat body. To the extent that the eicosanoid systems in insects are similar to the ones in mammals, studies on PLA, are of central importance. Moving this field beyond its infancy is a major desideratum.

8.1.3.2 Eicosanoid biosynthesis in tobacco hornworm tissues The idea

2o

"

1

14

1

I

0

1

0

20

10

TIME

PROTEIN (MG)

o ! 0

10

20

30

40

50

TEMPERATURE ('JC)

60

70

5

30

40

(MIN)

. , . , . , . , . , . 6

7

8

9

10

, 11

PH

FIG. 22 The effects of incubation conditions on PLA2 activity in microsomal-rich preparations from fat bodies of tobacco hornworm larvae. Each point is the mean of three or four experiments, and the error bars indicate one SE. As discussed in the text, these data indicate that a PLAz activity that can hydrolyse 20:4n-6 from phospholipids is present in fat body.

190


that eicosanoids mediate the cellular immune responses of an invertebrate animal opens a novel and very broad field for investigations. One of the more pressing questions relates to the identities of the active eicosanoids: which of many possible eicosanoids are biosynthesized by immune tissues? We are currently addressing this question in fat body and haemocytes from tobacco hornworms. A detailed report of the information presented here in outline form is forthcoming (Stanley-Samuelson and Ogg, 1994). All of the experiments were performed with fat body tissues harvested from second-day, fifth instar larvae of the tobacco hornworm. In our general protocol, microsomal-rich fractions were incubated with radioactive arachidonic acid. The reactions were stopped, and the eicosanoid products were extracted with acidified ethyl acetate. The reaction products were separated by 1-dimensional and 2-dimensional TLC, and the radioactivity in each fraction was assessed by liquid scintillation counting. The details of the protocol are similar to the methods of Howard et af. (1986). Here, it is useful to comment on specific techniques. The reactions were carried out in 1.0 ml total volume. The experiments were preceeded by a 3-min preincubation with all reaction components except the protein source. The preincubation brings all components of the reaction to the incubation temperature before the experiment begins. This is an important step because, compared with other studies of insect PG biosynthesis, we use far shorter incubation periods. After spotting the samples, the TLC plates were developed in solvent system A9 (Hurst et af., 1987), formed by mixing ethyl acetate:trimethyl pentane:distilled water:acetic acid (55:25:50:10, v/v) in a separatory funnel, then allowing the phases to separate. The lower aqueous layer was removed, and the organic layer was dried by addition of a few milligrams of sodium sulphate. Again, this is important because the chromatography is influenced by the presence of small amounts of water. Moreover, chromatography with this system differs from the usual procedures because the TLC plates are placed in the developing chamber immediately after the solvent system, without an equilibration period. The TLC plates require approximately 2.5 h to completely develop, after which the plates were air-dried. Fractions were visualized by exposure to iodine vapours. 8.1.3.2.1 Control experiments. PUFAs are labile to spontaneous reactions with oxygen, which can create reaction artefacts. We control for these possibilities by conducting “zero-time’’ reactions. In these control experiments, the substrate was pre-incubated in parallel with our biosynthesis experiments. At time zero, the pre-incubations are stopped. Extraction and analysis of the products follow the same procedures just described for the biosynthesis experiments. We include a zero-time reaction in every experiment. Values from these control experiments were used to correct the corresponding values in the prostaglandin biosynthesis experiments. The fat body preparations synthesize substantial amounts of four


191

prostaglandins, PGF2alpha,PGE2, PGD2 and PGA2. Under most conditions, PGA2 was the major product, although we will show that the incubation conditions alter the overall pattern of prostaglandin biosynthesis. The effects of altering our standard in vitro conditions are presented in the following paragraphs. Figure 23 shows the relationship between fat body microsomal protein concentration and prostaglandin biosynthesis. These data indicate that the optimal protein concentration for prostagladin biosynthesis was about 0.75-1.0 mg. These findings informed the use of 1.0 mg microsomal protein in subsequent experiments. These results were obtained from 10-min incubation periods, which we next show is not the optimal incubation period.

275 250 225

+-PGF2

200

-8- PGE2

175 c

1 .

+PGD2 +PGA2

150 125

100 75

50 25 0 0

1

2

3

6 Protein (mg)

4

5

7

8

9

10

FIG. 23 Eicosanoid biosynthesis by tobacco hornworm fat body preparations: the effect of microsomal-rich protein concentration. Each bar represents the mean of three experiments, and the error bars indicate one SE. These data show that PLAl is the major product that is biosynthesized under our conditions.


192

4000

3500

3000

E". 1 g

+PGF2 +PGE2

2500

uPGD2 +PGA2

2000

n

1500

1000 500 0 0

1

2

3

4

5

6

7

8

9

10

Time (min) FIG. 24 Eicosanoid biosynthesis by tobacco hornworm larvae fat body preparations: the effect of incubation time. Each bar represents the mean of three experiments, and the error bars indicate one SE. These data indicate that relatively short incubation periods, on the order of 1 min, are optimal in vitro PG biosynthesis

conditions. Figure 24 shows the profiles of prostaglandin biosynthesis at 10 incubation intervals from 0.1 to 10.0 min. The largest total prostaglandin biosynthesis (C. of all four products > 3400 pmol/mg protein) appeared in 1.0-min incubations. Total prostaglandin production was diminished in longer incubation periods, from about 2100 pmol/mg protein at 2 min to about 400 pmoVmg protein at 10 min. Similarly, total prostaglandin biosynthesis was reduced in very short incubation periods, those less than 1.0 min. Although PGA2 was the major product in most incubation periods (Fig. 24), the duration of the incubation appeared to alter the overall profiles of prostaglandin biosynthesis. At 0.1 min, the shortest incubation, PGFZalpha was the only detectable product. At 0.25 min, all four prostaglandins were formed. Again, PGFZalpha was the major product (> 1000 pmol/mg protein), while PGA2 biosynthesis was just detectable (c. 20 pmol/mg protein). At all incubation periods from 0.5 to 10 min, PGA2 was by far the dominant


193

product, by 3-10-fold. Substantial amounts of PGDZ were produced in all the incubation periods beyond 0.1 min; PGD2 synthesis varied with incubation duration from a high of about 700 pmoUmg protein at 0.25 min to below 75 pmol/mg protein after 4 min (Fig. 24). Biosynthesis of PGD2 appeared to be consistently greater than PGEz. PGE2 and PGFZalpha were also produced in substantial levels, however, the quantitative relationship between these two products was sensitive to the duration of incubations. For example, relative to PGE2, PGFZalphawas produced at 2.5-fold levels in 0.25-min incubations, and at 0.8-fold in 1.O-min experiments. All subsequent experiments featured 2-min incubations. The influence of incubation temperature on the profile of prostaglandin biosynthesis in shown in Fig. 25. Total biosynthesis increased from < 100 pmol/mg protein at 20°C to > 2100 pmol/mg protein at 30°C. Total biosynthesis sharply decreased at temperatures about 30"C, to about 1280 pmol/mg protein at 35°C and to < 150 pmoUmg protein at 40°C. Negligible biosynthesis occurred at 45°C.

1750 2ooo

'I

n

I

-F+

PGFP

+PGEP +PGD2 --t

PGA2

1000

750 500

i 20

/F\

TI

25

30

35

40

45

Temperature (C ) FIG. 25 Eicosanoid biosynthesis by tobacco hornworm larvae fat body preparations: the effect of incubation temperature. Each bar represents the mean of three experiments, and the error bars indicate one SE. These data emphasize the point that incubation conditions can influence the overall profile of PG biosynthesis.

194


The temperature effects recall our theme of the effects of incubation conditions on biosynthesis of individual components. At 25"C, PGD2 was the major product (> 400 pmol/mg protein compared to about 180 pmol/mg protein for PGA2), while at 30"C, biosynthesis of PGA2 was about an order of magnitude greater than of PGD2 (> 1700 compared to about 160 pmol/ mg protein). Although it was produced in lower abundance at the lower temperatures, PGFZalpha was the major prostaglandin, while PGA2 synthesis was reduced at 35°C. PGFZalphn biosynthesis appears to be quite temperature sensitive, because this product was not synthesized at 40 and 45°C. PGA2 biosynthesis is thought to proceed from PGH2 through PGE2, which can be converted to PGAz and PGB2; PGA2 is not directly formed from PGHz in mammals. The relatively high levels of PGA2 biosynthesis by fat body preparations could be an artefact of spontaneous rearrangements of PGE2 during our analytic steps, a product of nearly quantitative PGE2 metabolism, or a product of an unknown biosynthetic route that has not yet come to light. We investigated these possibilities by incubating fat body preparations with radioactive PGE2 under our standard experimental conditions. Products were extracted from the reaction mixtures and separating according to our usual chromatographic procedures. About 90% of the radioactivity was recovered in a fraction that co-chromatographed with authentic PGE2, while 3%, 4% and 2.6% was recovered as PGFZalphar PGD2 and PGA2, respectively. These data indicate that PGE2 is not quantitatively converted to PGA2, and that the analytical procedures did not produce PGA2 as an artefact. Our working hypothesis is that the hornworm fat body produces PGAz by way of an unidentified intermediate. The fat body preparation is sensitive to two non-steroidal antiinflammatory drugs, indomethacin (not shown) and naproxin (Fig. 26). Addition of 1.O VM indomethacin reduced PGA2 biosynthesis from over 1700 pmol/mg protein to about 58 pmol/mg protein in our standard conditions. Biosynthesis of PGD2, PGE, and PGFZalphawas similarly sensitive to indomethacin, although on a smaller scale. PGFZalphabiosynthesis was reduced from about 150 pmol/mg protein to about 7 pmol; PGEz from about 115 pmol to 13 pmol, and PGD2 from > 160 pmol to 35 pmoV mg protein. Prostaglandin biosynthesis was also inhibited by low dosages of naproxin (Fig. 26). These data show considerable sensitivity to naproxin (ICs0 for naproxin is 0.7 VM). Indomethacin and naproxin appeared to increase the biosynthesis of one or more lipoxygenase products (Fig. 26). Standard separation of radioactive products on TLC yielded two new products, one of which co-chromatographed with authentic 15-HETE. The TLC solvent system Aq is unlikely to distinguish among a number of possible HETEs, and we take this finding to indicate that inhibition of cyclooxygenase activity leaves substrate available for lipoxygenases that remain to be identified. 8.1.3.2.2 The fat body preparation requires co-factors used in mammalian

195


f

g

PGA2

3000

S! 15-HETE

' '

5. 2500 2000 1500

1000 500

0 0.0

0.00001 0.0001

0.001

0.1

1.0

mM Naproxin FIG. 26 Eicosanoid biosynthesis by tobacco hornworm larvae fat body preparations: the effect of the cyclooxygenase inhibitor naproxin. Each bar represents the mean of three experiments, and the error b a k indicate one SE. Cyclooxygenase inhibitors can produce an apparent stimulation of lipoxygenase activity, as shown. It is generally thought that the increased lipoxygenase activity results from the increased substrate availability due to inhibiting cyclooxygenase activity.

preparations. PG biosynthesis has been investigated in a number of insect and other invertebrate systems (Stanley-Samuelson and Loher, 1986; Stanley-Samuelson, 1991). The general procedures usually include incubating a tissue homogenate with radioactive substrate, then monitoring eicosanoid biosynthesis by a form of radio-chromatography . Based on experience with mammalian preparations, most experiments with insects were performed in a buffer amended with a cocktail of co-factors, usually a mixture of reduced glutathione and haematin. Haemoglobin or myoglobin is often used in place of haematin. The roles of these co-factors relate to the mechanism of cyclooxygenase activity. In mammals, PGH synthase is a combination of two activities, a cyclooxygenase and a hydroperoxidase, in a single holoenzyme molecule (Fig. 5 ) . Cyclooxygenase activity converts a PUFA to PGG. Reduced glutathione is oxidized by the attendant hydroperoxidase in the process of reducing PGG to PGH (Kulmacz and

196


Lands, 1987). The cyclooxygenase and hydroperoxidase activities require haeme, specifically iron protoporphyrin IX (Hemler et al., 1976). The globins myoglobin and haemoglobin also stimulate both activities of PGH synthase in vitro, although the stimulation is due to release of haeme from the globins. If the PG biosynthetic enzymes in insects are similar to their counterparts in mammals, we would expect that the insect preparations would similarly require haeme and reduced glutathone. These co-factors are included in incubations with insect preparations; however, I do not know if they are required for eicosanoid biosynthesis. We used the fat body preparation to test the idea directly, and found that PG biosynthesis was reduced to zero or nearly zero in the absence of the co-factor cocktail (Stanley-Samuelson and Ogg, 1994). We do not yet know if the haeme binds to the insect and mammalian cyclooxygenases in the same way. Nevertheless, we conclude that the insect PGH synthase is similar to its mammalian counterpart on the requirement for co-factors. We are in the early stages of similar experiments with haemocytes. In these experiments, the cells are pelleted from haemolymph by gentle centrifugation. The cells are taken up in buffer, then homogenized by sonication. Microsomal-rich fractions are prepared as described for the fat body experiments. These haemocyte preparations also convert radioactive 20:4n-6 into eicosanoids, mainly one or more lipoxygenase products. The products co-chromatograph with authentic 15-HETE on 1- and 2-dimensional TLC, but they have not been characterized to satisfaction. These results take their significance with respect to the effects of eicosanoid biosynthesis inhibitors on cellular immune responses. Let us explore future directions by looking at the salient shortcomings in the knowledge that has accrued so far. Our original work indicated that inhibition of eicosanoid biosynthesis compromised cellular immunity (StanleySamuelson et al., 1991). We documented the presence of eicosanoidprecursor PUFAs in haemocytes (Ogg et al., 1991) and fat body (Ogg and Stanley-Samuelson, 1992). The work just described shows the presence of a cellular PLA2 activity in fat body, and that fat body and haemocytes are able to convert radioactive 20:4n-6 into eicosanoids. It would appear, in broad view, that the major elements of the eicosanoid system are in place. What additional information do we need to rigorously test the hypothesis that eicosanoids mediate cellular immune responses in insects? First, we have not shown that our treatments with pharmaceutical inhibitors of eicosanoid biosynthesis quantitatively reduced eicosanoid titres in fat body, haemocytes or haemolymph. Second, we have no information on quantitative changes in eicosanoid biosynthesis or titres following infection. Third, we do not know whether or not bacterial infection impacts the pattern of eicosanoid biosynthesis in the relevant tissues. Fourth, we lack information on the specific cellular events that eicosanoids may influence. Information on these


197

points will be useful in testing the eicosanoid hypothesis and in gaining an understanding of the specific roles of eicosanoids in immunity. Our current work on biosynthesis is an essential prerequisite for quantitative studies because the appropriate techniques utilize antibodies to specific eicosanoids. 8.1.3.3 The molecular biology of eicosanoids I offer a few words on the molecular biology of eicosanoids in this final section on desiderata. These comments are not meant to pose as a review of this rapidly growing literature; contrarily, the aim is to recognize that molecular methodologies are among the many technical tools that have become available. Advances in molecular biology have yielded detailed insights into mammalian eicosanoid systems. Genes for PG synthases and 5-, 12-, and 15lipoxygenases have been cloned from several mammalian species, including humans. Genes for other proteins in mammalian eicosanoid systems, including genes for eicosanoid receptor proteins and for PG-catabolic enzymes have also been cloned. These cloning efforts facilitated more penetrating studies of regulation of eicosanoid systems. Drawing on the recent volume edited by Bailey (1991), we now have insights into transcriptional regulation of a mouse PGH synthase gene, regulation of cyclooxygenase gene expression in vascular endothelial cells, and the mode of action of glucocorticosteriods. We also have information on the genes for PG-dehydrogenase, an enzyme that serves to catabolize biologically active PGs into inactive forms. The chromosomal localization, structure and expression of 12-lipoxygenase genes in human platelets and umbilical vein endothelial cells was recently described by Funk et al. (1992). Similarly, the structure of the gene for PGD synthase in rat brain was described by Igarashi et al. (1992). The growing wealth of information on the sequences and structures the genes associated with the eicosanoid systems in mammals may help us design probes for analogous systems in insects and other invertebrates. Progress in this area of invertebrate eicosanoid systems may very effectively help move toward a more complete understanding of the biological significance of eicosanoids in insect physiology.

8.2

COMPARATIVE PHYSIOLOGY OF EICOSANOIDS

The biological significance of eicosanoids transcends taxonomic boundaries. Our knowledge of the actions of eicosanoids in vertebrate and invertebrate systems is rapidly expanding. The new information is making it clear that similar eicosanoid actions occur in a broad range of animal phyla. A good example of this point is regulation of ion and water transport. We have seen that eicosanoids are involved in mammalian kidney function, in bladder and skin transport in amphibians, in epithelial transport in marine and freshwater molluscs, and most recently in mosquito Malpighian tubules. The

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unifying conceptualization is that at the cellular level of organization, eicosanoids may regulate fundamental cellular events in similar cells in animals from quite diverse phyla. To the extent that this generalization holds, it is reasonable to postulate that some eicosanoid actions may be hypothesized from narrow examples. Building on the realization that eicosanoids impact on ion transport in a wide range of animal cells, it is not unreasonable to predict that these molecules may be important in the physiology of Malpighian tubules, sensu latu, as well as other transporting tissues such as the insect rectum. Similarly, we now know that eicosanoids are involved in the cellular immunity of mammals, lower vertebrates and tobacco hornworm larvae. I suppose that eicosanoids regulate actions in the immunocytes of many, if not all, animals. Indeed, Barge and Karp (personal communication) found that inhibition of eicosanoid biosynthesis in cockroaches similarly compromised cellular immunity. Examples of eicosanoid actions that may be fundamental in animal cells abound. To touch upon two, I recently reviewed the roles of eicosanoids in prevention of polyspermic fertilizations in sea urchins eggs (StanleySamuelson, 1991). Also, we have just seen that eicosanoids are important in synaptic physiology. Eicosanoid roles in basic cellular actions suggest that these molecules are elements in the physiology of all animals. One implication of this view is that insights from one animal system may shed light on the regulatory physiology of other, distantly related, systems. There are eicosanoid actions that do not appear in a wide range of animal phyla. An important example comes from the first discovery of a physiological action of PGs. In studies of the reproductive biology of. mammals, it was found that seminal fluids and extracts of prostate glands stimulate contractions in uterine smooth muscle (Kurzrok and Lieb, 1930; Goldblatt, 1933). It is now known that various PGs induce contractions or relaxations in smooth muscle from many sources. One might reasonably expect that PGs similarly produce contractions of visceral muscles of insects. Observations on the oviducal muscles of the field cricket T. commodus by Loher (personal communication) indicate that PGs do not stimulate muscle contraction. Similarly, Cook et al. (1984) observed that biogenic amines, but not PGE2, cause contraction of the oviduct musculature of the cockroach Leucophaea maderae. Such similarities and differences in eicosanoid biology among animal taxa are the grist of comparative physiology. Comparative views contribute to our deeper understanding of physiology by placing apparently isolated findings onto a broader landscape. These broader perspectives can serve also as a rough guide, when taken with due scepticism, to areas of physiology that would be considerably enhanced by an understanding of the biochemistry and physiology of eicosanoids. Inquiry along these lines will produce rapid and profound advances in insect physiology.


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Acknowledgements

I am grateful to three colleagues who devoted their time and energy to reading and making useful comments on an earlier draft of this essay: Drs Ralph W. Howard, David H. Petzel and John E. Foster. Thanks, also, to Mr Ken Jensen for creating the figures. This chapter is paper number 9849, in the Nebraska Agricultural Research Division Journal series, and Contribution number 792 of the Department of Entomology, University of Nebraska-Lincoln. Investigations in my laboratory are supported by the University of Nebraska Agricultural Research Division, the Department of Entomology and by NIH grant AI31509.

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Tinoco, J. (1982). Dietary requirements and functions of alpha-linolenic acid in animals. Prog. Lipid Res. 21, 1-45. Tinoco, J., Babcock, R., Hincenbergs, I., Medwadowski, B. and Miljanich, P. (1979). Linolenic acid deficiency. Lipids 14, 166-173. Tobe, S. S. and Loher, W. (1983). Properties of the prostaglandin synthetase complex in the cricket Teleogryllus commodus. Insect Biochem. 13, 137-141. Toolson, E . C. (1987). Water profligacy as an adaptation to hot deserts: water loss rates and evaporative cooling in the Sonoran Desert cicada, Diceroprocta apache (Homoptera: Cicadidae). Physiol. Zool. 60, 379-385. Toolson, E. C. and Hadley, N. F. (1987). Energy-dependent facilitation of transcuticular water flux contributes to evaporative cooling in the Sonoran Desert cicada, Diceroprocta apache (Homoptera: Cicadidae). J . exp. Biol. 131, 439444. Turunen, S. (1973). Utilization of fatty acids in Pieris brassicae reared on artificial and natural diets. J . Insect Physiol. 19, 1999-2009. Turunen, S. (1974). Polyunsaturated fatty acids in the nutrition of Pieris brassicae (Lepidoptera). Ann. Zool. Fenn. 11, 300-303. Turunen, S. and Parnanen, S. (1987). Eicosapentaenoic acid: incorporation into phosphatidylinositol and other lipids in Pieris brassicae. Insect Biochem. 17, 891-895. Uchida, M., Izawa, Y. and Sugimoto, T. (1987). Inhibition of prostaglandin biosynthesis and oviposition by an insect growth regulator, Buprofezin. in Nilaparvata lugens Stal. Pest. Biochem. Physiol. 27, 71-75. Uschian, J. M. and Stanley-Samuelson, D. W. (1993). Phospholipase A2 activity in the fat body of the tobacco hornworm Manduca sexta. Arch. Insect Biochem. Physiol. 24, in press. Vance, D. E. and Vance, J . E. (Eds) (1985). “Biochemistry of Lipids and Membranes”. BenjaminKummings, Menlo Park, Calif. Vance, D. E. and Vance, J. (Eds) (1991). “Biochemistry of Lipids, Lipoproteins and Membranes”. Elsevier, Amsterdam. Wakayama, E. J., Dillwith, J. W:and Blomquist, G. J. (1985). Occurrence and metabolism of arachidonic acid in the housefly, Musa domestica (L.). Insect Biochem. 15, 367-374. Wakayarna, E. J., Dillwith, J. W. and Blomquist, G. J. (1986a). Occurrence and metabolism of prostaglandins in the housefly, Musa dornestica (L.). Insect Biochem. 16, 895-902. Wakayama, E. J., Dillwith, J. W. and Blomquist, G. J. (1986b). Characterization of prostaglandin synthesis in the housefly, Musca domestica (L.). Insect Biochem. 16. 903-909. Weinheimer, A. J. and Spraggins, R. L. (1969). The occurrence of two new prostaglandin dervatives (15-epi-PGA2 and its acetate methyl ester) in the gorgonian Plexaura homomalla: chemistry of coelentrates IV. Tetrahedron Lett. 59, 5185-5188. Whittier, J. M. and Crews, D. (1989). Mating increases plasma levels of prostaglandin FZalpha in female garter snakes. Prostaglandins 37, 359-366. Witters, N. A. (1991). “Immunohistochemica1 Localization of Eicosanoids in Arthropod Tissue: Implications for Invertebrates”. M.S. Thesis, University of Nebraska, Lincoln. Wlodawer, P. and Baranska, J. (1965). Lipolytic activity of the fat body of the waxmoth larvae. Acta Biochim. Polonica 12, 25-36. Yamaja Setty, B. N. and Ramaiah, T. R. (1979). Isolation and identification of prostaglandins from the reproductive organs of male silkrnoth, Bombyx mori ( L . ) . Insect Biochem. 9, 613-617.

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Yamaja Setty, B. N. and Ramaiah, T. R. (1980). Effect of prostaglandins and inhibitors of prostaglandin biosynthesis on oviposition in the silkmoth Bombyx mori. Indian J. exp. Biol. 18, 539-541. Yamaja Setty, B. N. and Ramaiah, T. R. (1982). Effect of PGE, on lipid mobilization from pupal fat bodies of the silkworm, Bombyx mori. Indian J . exp. Biol. 19, 115-118. Yorio, T., Page, R. D. and Frazier, L. W. (1991). Prostaglandin regulation of H+ secretion in amphibian epithelia. Am. .I. Physiol. 260 (Regulatory Integrative Comp. Physiol. 29), R866R872. Zinkler, D. (1975). Zum Lipidmuster der Photorezeptoren von Insecten. Verh. dt. 2001. Ges. 1974, 28-32.

Cellular and Molecular Actions of Juvenile Hormone 1. General Considerations and Premet a m o r phic Actions Lynn M. Riddiford Department of Zoology, University of Washington, Seattle, Washington 98195, USA

Introduction 213 1.1 Biological roles of juvenile hormone 215 1.2 Current concepts of hormone action 219 Embryonic actions of juvenile hormone 224 Premetamorphic actions of juvenile hormone 225 3.1 Regulation of cellular commitment by juvenile hormone 225 3.2 Epidermis 226 3.3 Fat body 235 3.4 Muscle 239 3.5 Nervous system 242 3.6 Other morphogenetic actions 233 Mechanism of JH action 244 4.1 Juvenile hormone binding proteins in the haemolymph 246 4.2 Intercellular juvenile hormone receptors 247 4.3 Morphogenetic action of JH: modulation of ecdysteroid action 251 Juvenile hormone analogues as insect growth regulators 253 Conclusions 254 Acknowledgements 255 References 256

Introduction

Since its discovery by Wigglesworth (1934, 1936), juvenile hormone (JH) has been known to be critical for the regulation of both metamorphosis and ~~

I dedicate this review to the memory of Carroll M. Williams (1916-1991) who first introduced me to the fascination of juvenile hormone and its role in metamorphosis. His serendipitous discovery of a natural repository for juvenile hormone enabled its later chemical identification. Moreover, his insightful studies of the biological actions of juvenile hormone helped to provide the basis for the current investigations of its cellular and molecular actions as well as for the use of juvenile hormone analogues in insect control. He was an outstanding insect physiologist and endocrinologist whom the field sorely misses. ADVANCES I N INSECT PHYSIOLOGY VOL ?? ISBN Wl2-034224')

L. M. RlDDlFORD

214

JH I1

(a)

4-MeJH I

Methyl Famesoate 0

methoprene

AAAAAAA 0

hydroprene NHCO

fenoxycarb

(b)

pyriproxyfen

0

ewo3 N

FIG. 1 (a) Natural juvenile hormones found in insects. The stereochemistry is shown for those for which it has been determined. JHB3, JH 111 bisepoxide. (b) Juvenile hormone analogues that are used as insect growth regulators. (Fig. l a is courtesy of Dr David Schooley (unpublished).) reproduction in insects. Although initially thought to be released by the brain, Bounhiol (1938) and Wigglesworth (1940) later showed that it was secreted by the corpora allata, a pair of glands in the head just posterior to the brain. Over the years since its discovery, JH has been isolated (Williams, 1956) and chemically identified as a sesquiterpenoid (Roller et al., 1967). We now know that there are different forms of the secreted hormone, differing by side-chain length or by epoxidation: JH 0, I, and I1 being found in Lepidoptera with JH I11 in most other insects (Schooley et al., 1984)

CELLULAR AND MOLECULAR ACTIONS OF JUVENILE HORMONE

215

(except for J H I in the hemipteran Riptortus clavatus (Numata et al., 1992)), JH I11 bisepoxide in higher Diptera (Richard et al., 1989b; Cusson et a f . , 1991), and small amounts of methyl farnesoate, the immediate precursor of J H 111, in cockroach embryos and larvae (Bruning et al. 1985; Cusson et al., 1991) (Fig. l a ) . The action of these natural hormones can be mimicked by a number of small lipophilic molecules such as seen in Fig. 1b. Although much is known about the biological actions of these relatively small, lipophilic molecules on the organismal level, much less is understood about their actions on the cellular and molecular level (see also Kumaran, 1990). In this two part review, we will focus on the latter. After a brief review of the biological role of J H and of the prevailing concepts of the modes of actions of hormones, this first part will concentrate on the premetamorphic actions of J H and our present understanding of the molecular basis of this action. Part I1 in the succeeding volume (Wyatt and Davey , unpublished) will concentrate on the action of JH in reproductive maturation in the adult.

1.1

BIOLOGICAL ROLES OF JUVENILE HORMONE

1.1.1 Premetamorphic Juvenile hormone is first produced by the corpora allata in the late embryo and appears to be important for normal dorsal closure, formation of the larval cuticle, and differentiation of the midgut (Bergot et af., 1981; Dorn, 1982; Bruning and Lanzrein, 1987). The corpora allata remain active throughout larval life until the final instar although there may be fluctuations in activity depending on the stage (Tobe and Stay, 1985). There also are fluctuations in the activity of the main degradative enzyme, JH esterase, in the haemolymph with a peak before each larval ecdysis (Jones and Click. 1987; Jesudason et al., 1990; Roe and Venkatesh, 1990). The result is that J H is generally present in the larval haemolymph during both the feeding intermoult phase and the moulting phase (Fain and Riddiford, 1975; Lanzrein et al., 198.5; Grossniklaus-Burgin and Lanzrein, 1990). The role of J H during the feeding phase is uncertain although it seems to be necessary for the maintenance of larval-specific organs such as the crochets (hooked setae) on the abdominal prolegs of caterpillars (Fain and Riddiford, 1977), and possibly for the continued high production of larval pigment proteins (Goodman et al., 1987). It also likely promotes the high metabolism associated with growth (Sehnal and Slama, 1966; Slima and Kryspin-Sorenson, 1979; Steele, 198.5; Stephen et al., 1988), and in the holometabolous insects, may be essential €or the continued proliferation of the imaginal discs (Davis and Shearn, 1977; Oberlander, 1985). This intermoult JH could also be important in behaviour associated with feeding

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since starved lepidopteran larvae show increased locomotory behaviour (0. Dominick, personal communication) and a greatly increased J H titre (Cymborowski et al., 1982). Finally, at least in some Lepidoptera, J H seems to be important for the production of the prothoracicotropic hormone (PTTH) in the penultimate instar larva (Hiruma, 1986). The presence of J H at the onset of the ecdysteroid rise that causes each moult is critical to prevent metamorphosis as first shown by Wigglesworth (1934) in the blood-sucking bug, Rhodnius prolixus, and later by Bounhiol (1938) in the silkworm, Bombyx mori. Thus, during a larval moult initiated in the presence of JH, ecdysteroid causes the production of a new larval cuticle and any changes in the viscera associated with the moult. In the absence of JH at this critical time, the ecdysteroid causes major changes in all the tissues which result in transformation of the hemimetabolous larva into the adult or of the holometabolous larva into the pupa. Later during the course of the larval moult, J H may also affect the pigmentation of the new cuticle that will be formed as well as pigmentation of the epidermis itself (Nijhout and Wheeler, 1982; Riddiford, 1985; Goodman et al., 1987; Riddiford and Hiruma, 1988). During the final larval instar the titre of J H differs in hemimetabolous and holometabolous insects. In hemimetabolous insects such as the cockroach Nauphoeta cinerea (Lanzrein et al., 1985; Fig. 2a), the cricket Teleogryllus commodus (Loher et al., 1983), and Locusta migratoria (HuibregtseMinderhoud et al., 1980), J H levels are low to undetectable early in the final instar, and there are one or several small rises of ecdysteroids that precede the moulting surge. Such ecdysteroid rises in Locusta migratoria (Hirn et al., 1979) in the absence of J H (Huibregtse-Minderhoud et al., 1980) initiate metamorphic changes such as mitosis in the wing pads (Brehelin and Aubry, 1982), the development of the flight muscles (Van den Hondel-Franken, 1982), and the attainment of differentiative competence in male accessory glands (Gallois, 1989). At the onset of the main rise of ecdysteroid that initiates the moult, there is a critical period during which J H must be absent to allow the metamorphic changes in the epidermis so that the adult cuticle and pigments may be formed such as in the hemipterans, Oncopeltus fasciatus and Rhodnius prolixus (Smith and Nijhout, 1981; Nijhout, 1983). Changes in the fat body at metamorphosis that cause altered responsiveness to J H of genes such as the vitellogenins (Dhadialla and Wyatt, 1983) and the persistent storage protein (Wyatt et al., 1992a) in Locusta are presumably similarly determined. In holometabolous insects such as Lepidoptera (Granger et al., 1982; Janzen et al., 1991) and Diptera (Richard et al., 1989a), J H is still being produced by the corpora allata in the early part of the final instar. Then synthesis declines, and, in the case of the tobacco hornworm, Manduca sexta, and other lepidopterans, the methyl transferase disappears so that J H acid instead of J H is released (Bhaskaran et al., 1986). This cessation of

217


(DAYS)

I

K W

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r',

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72

48

Is'

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E

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.

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FIG. 2 Schematic of juvenile hormone (dotted line) and ecdysteroid (solid line) titres. (a) Nauphoeta cinerea (data from Lanzrein et al., 1985); (b) Manduca sexta (data from Fain and Riddiford, 1975; Bollenbacher et al., 1981; Curtis et al., 1984; Wolfgang and Riddiford, 1986; Baker et a [ . , 1987); (c) Drosophila melanogaster (data from Kraminsky et al., 1980; Handler, 1982; Sliter et al., 1987; Bownes and Rembold, 1987). E, ecdysis; H , hatch; P, pupariation; W, wandering.

synthesis combined with a greatly increased J H esterase activity in the haemolymph (Roe and Venkatesh, 1990) causes a decline of the JH titre to undetectable levels within the first third of the instar (Baker et a f . , 1987; Grossniklaus-Burgin and Lanzrein, 1990) (Fig. 2b). In Drosophila melanogaster the titre drops early in the 3rd larval instar with a brief upsurge just prior to pupariation (Sliter et al., 1987; Bownes and Rembold, 1987) (Fig. 2c). In this case, early in the instar, the J H epoxide hydrase is high; whereas

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after pupariation, the main metabolizing enzyme is the J H esterase (Campbell et al., 1992). The corpora allata in wandering final instar Drosophifu larvae make primarily J H I11 bisepoxide (Richard et al., 1989b). Whether they have shifted from production of J H I11 during the larval intermoults has not been determined. In final instar Manduca larvae, this drop in J H allows the brain to release PTTH (Rountree and Bollenbacher, 1986) and thus a small rise in ecdysteroid is seen on the final day of feeding (Bollenbacher el a f . , 1981; Wolfgang and Riddiford, 1986; Gilbert, 1989) (Fig. 2b). This ecdysteroid in the absence of J H acts on the tissues to initiate metamorphosis; on the epidermis to cause it to become pupally committed so that it can no longer make larval products even when exposed to a larval moulting environment (Riddiford, 1976); on the nervous system to cause the cessation of feeding and the onset of wandering behaviour (Dominick and Truman, 1985) and commitment to metamorphosis (Weeks and Levine, 1990); and on the fat body to cause a switch from the nutritive and synthesis mode to the storage mode (Dean et al., 1985; Webb and Riddiford, 1988a, b). After the larva has found a suitable place to pupate, PTTH is again released, and ecdysteroid rises in the presence of J H (Bollenbacher et a f . , 1981; Baker et al., 1987) to cause the formation of the pupa. This J H prevents adult differentiation of imaginal discs and other imaginal precursors (Williams, 1961; Kiguchi and Riddiford, 1978). Just before pupal ecdysis the J H esterase again transiently increases (Jesudason et a f . , 1990) so that the new pupa has no JH. Consequently, the subsequent ecdysteroid rise (Warren and Gilbert, 1986) in the pupa occurs in the absence of JH to cause the formation of the adult (Williams, 1961).

1.1.2 Adult

In the adult, J H is again used by most insects for regulation of various aspects of reproductive maturation and behaviour (Koeppe et al., 1985; Bownes, 1986; Wyatt, 1991). The precise nature of its role varies with the insect and its particular reproductive strategy. These actions of J H will be covered in detail in Part I1 (Wyatt and Davey, unpublished) and just briefly summarized below. In feeding female insects, J H generally controls one or several aspects of oogenesis and/or vitellogenesis: (1) previtellogenic actions on the developing fat body and/or oocyte such as stimulation of ribosomal proliferation in fat body prerequisite for vitellogenin synthesis and the development of the pinocytotic complex in mosquitoes necessary for vitellogenin uptake (Engelmann, 1987; Wyatt, 1988; Raikhel and Dhadialla, 1992); (2) vitellogenin synthesis in the fat body as in cockroaches (Engelmann, 1987) and locusts (Wyatt, 1988); ( 3 ) vitellogenin uptake into the oocyte as in


219

Rhodnius (Davey, 1981); (4) later phases of oocyte maturation such as an increased rate of vitellogenesis followed by hydration of the egg in Manduca (Nijhout and Riddiford, 1979). JH may also control the synthetic activities of the accessory glands such as the production of the oothecins that form the egg case in cockroaches (Scharrer, 1946; Pau et al., 1986). In some cases, sex pheromone production is also controlled by J H (Cusson and McNeil, 1989). In males gonad maturation occurs as part of the metamorphic process and therefore is under the control of ecdysteroid acting in the absence of JH (Happ, 1992). Initial accessory gland differentiation is also regulated similarly. In the adult, JH may be necessary for the continued production of the accessory gland secretions. Male reproductive behaviour may also be regulated by JH such as the courtship behaviour of Gomphocerus rufus (Loher and Huber, 1966).

1.2

CURRENT CONCEPTS OF HORMONE ACTION

Hormones are classically defined as molecules secreted by an endocrine organ or a neuron that circulate in the blood and act on distant targets. The target may be any cell that has specific receptors for the hormone. Hormones may be broadly classified into two groups: (1) steroid and other lipophilic hormones which cross the cell membrane and bind to intracellular receptors; and (2) protein, peptide and catecholamine hormones which bind to specific membrane receptors and modulate intracellular actions via second messengers. 1.2.1 Steroid hormone action Since the pioneering work of Clever and Karlson (1960) showing that ecdysone caused a rapid change in the puffing pattern of the salivary gland chromosomes of the midge, Chironomus tentuns, steroid hormones have been thought to enter cells, bind to intracellular receptors, then act directly on the genome. In the absence of hormone, these receptors are now known to be localized either in the cytoplasm (e.g. glucocorticoid receptors) or in the nucleus (e.g. oestrogen or thyroid hormone receptors) (Beato, 1989; Carson-Jurica et al., 1990). In the presence of the hormone, they all are found in the nucleus. The steroid hormone receptors all belong to a superfamily of proteins that bind to specific DNA sequences (“hormone response elements”) usually as dimers and either activate or inactivate the associated genes (Evans, 1988; Carson-Jurica et al., 1990; Laudet et al., 1992). The 66 amino acid DNA binding region of these proteins is most conserved within the family and includes two “zinc fingers” with each zinc coordinately bonded by four cysteine residues (Fig. 3). Three amino acids in the N-terminal finger of the

220

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FIG. 3 Diagram of the steroid hormone receptor protein. The enlargement of the C (DNA binding) region shows the sequence of the ecdysteroid receptor (Koelle et al., 1991) in the predicted two “zinc fingers” based on the known structure of the oestrogen receptor (Schwabe and Rhodes, 1991).

estrogen and the glucocorticoid receptors actually bind to the DNA, and the C-terminal finger participates in dimer formation and may interact with other nuclear proteins (Schwabe and Rhodes, 1991). The hormone binds to the C-terminal ligand-binding region of the receptor, induces translocation to the nucleus in cases where the unliganded receptor is in the cytosol, and causes DNA binding of the hormone-receptor complex. The C-terminal domain also contains sequences that are important in dimerization and in the transactivation functions of the hormone-receptor complex (Glass et af., 1990; Danielian et af., 1992). The N-terminal region of the receptor is critical also for the transactivation and may differ among the different isoforms of the receptor generated by alternative splicing or alternative promoters or by different genes (Carson-Jurica et af., 1990; Danielian et af., 1992). The steroid hormone-receptor complex may either activate or inactivate specific genes that have the appropriate hormone response elements (Wahli and Martinez, 1991; Lucas and Granner, 1992). Also, certain of these steroid hormone receptors may in the absence of hormone suppress the genes that they activate in the presence of hormone (thyroid hormone (Damm et al., 1989; Baniahmad er af., 1992); ecdysteroid (Cherbas et af., 1991; Dobens et al., 1991)). Usually they bind as homodimers to palindromic sequences of two 5-6 bp half-sites separated by variable spacing (Schwabe and Rhodes, 1991) although direct repeats have also been found (Lucas and Granner, 1992). Heterodimers are also found such as the


22 1

interactions of retinoid X receptors with both thyroid hormone and retinoic acid receptors to enhance the latter’s transcriptional activity (Zhang et al., 1992; Kliewer et al., 1992). Heterodimers can also be formed with other types of transcription factors at a “composite” response element (Miner and Yamamoto, 1991; Schule and Evans, 1991; Lucas and Granner, 1992); this greatly increases the potential for interactions among several signalling pathways within the cell. In addition to co-operativity in DNA binding, these different types of factors may interact at the protein level to change the nature of the transcription complex or may compete with one another for DNA binding. The steroid hormone-receptor complex also alters chromatin structure as indicated by changes in DNase hypersensitive sites (Rories and Spelsberg, 1989). Additionally, steroid hormones may regulate stability of particular mRNAs (Nielsen and Shapiro, 1990). Yet McKenzie and Knowland (1990) have shown that the stabilizing effect of oestrogen on vitellogenin mRNA in chick liver is likely due to pharmacological doses of the hormone, thus calling into question this effect. 1.2.2 Ecdysteroid action Studies on steroid-induced puffing patterns of the giant salivary chromosomes of Chironomus tentans (Clever and Karlson, 1960; Clever, 1964) and Drosophila melanogaster (summarized in Ashburner et al., 1974) were the first to show that a steroid could act directly on genes to initiate a cascade of gene activation and inactivation.. These studies led to a model for ecdysteroid action (Ashburner et al., 1974) in which the ecdysteroid coupled to the ecdysteroid receptor (EcR) acts differentially to regulate several classes of target genes. “Early” genes are turned on directly by the ecdysteroid-receptor complex whereas ‘‘late’’ genes are repressed by it. The early genes were thought to code for one or more regulatory proteins that induced a secondary response by repressing the early genes and activating the late genes. Recently, the Drosophila EcR has been cloned and shown to be a member of the steroid receptor superfamily (Koelle et al., 1991). Interestingly, EcR forms heterodimers with the ultraspiracle protein, another member of the same family, and together they bind both radiolabelled ecdysteroids and DNA sequences (Yao et al., 1992; Thomas et al., 1993) that contain a known ecdysone response element (Cherbas et al., 1991; Dobens et al., 1991; Luo et al., 1991; Ozyhar et al., 1991). Whether EcR forms heterodimers with other members of the steroid hormone receptor superfamily is not known. Immunocytochemical analysis shows that the EcR binds to both early and late chromosomal puff sites as postulated in the Ashburner model (W. S. Talbot and D. S. Hogness, unpublished as cited in Koelle et al., 1991).

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There are at least three different isoforms of the Drosophila EcR that share common DNA and ligand binding domains but differ in their Ntermini (Talbot et al., 1993). These isoforms show tissue- and stagespecificity in both quality and quantity. For instance, within the Drosophila nervous system, one form predominates at the onset of metamorphosis (pupariation), then disappears followed by the appearance of either the same or a different form. Both the isoform and the timing of its appearance during pupal and adult development are correlated with cell fate (Truman et al., 1993). The significance of the dynamics of these changes in receptor type during development is not yet understood. Several of the early, ecdysteroid-responsive genes in Drosophila, E75 (Segraves and Hogness, 1990), E74 (Burtis et al., 1990; Thummel et al., 1990) and the Broad complex (BR-C) (DiBello et al., 1991), have been found to encode multiple DNA binding proteins (see Andres and Thummel, 1992, for a review). Similar ecdysteroid-induced proteins have also been found in Lepidoptera (Palli et al., 1992; Jindra et al., 1992; Segraves and Woldin, 1992). Most of these mRNAs are induced at each moult and so do not seem to be stage-specific. Yet the BR-C mRNAs first appear at metamorphosis (Guay and Guild, 1991; Andres and Thummel, 1992), indicating that some may be stage-specific. These DNA-binding proteins are thought to regulate the secondary response to ecdysteroid by acting as transcription factors. Since the different forms are generated by use of alternative promoters or alternative splicing, the timing of the appearance of a particular form depends on its transcript length and on the sensitivity of its promoter to ecdysteroid (Thummel et al., 1990; Karim and Thummel, 1991, 1992). These transcription factors are not tissue-specific (Boyd et al., 1991) although their temporal appearance may differ in various tissues and even within cells of a particular tissue. For instance, in the 5th abdominal segment of Manduca 4th instar larvae, an ecdysteroid-induced member of the steroid hormone receptor superfamily, MHR3 (Palli et al., 1992), appears first in the epidermis that forms the crochets on the prolegs, then in the central intrasegmental regions, and finally in the black stripe region over a period of 12-15 h during the rise in ecdysteroid for the moult to the 5th instar (Langelan, Hiruma, Palli and Riddiford, unpublished). This pattern of appearance mimics that of the ability to moult after isolation from the prothoracic glands (Truman et al., 1974) except that it is delayed by about 6-8 h. Presumably it reflects the number of ecdysteroid receptors present in the various cells at the time of the onset of the ecdysteroid rise. The lack of stage- and tissue-specificity of these factors suggests that they likely interact combinatorially with other stage- and tissue-specific factors as well as with the EcRs to activate or inactivate the target genes in a cascading fashion.


223

1.2.3 Peptide hormone action Since peptides, proteins, and catecholamines cannot cross the cell membrane, they have transmembrane receptors that initiate intracellular actions. The largest class of these receptors are those that upon binding with the hormone activate a guanyl nucleotide binding protein (G protein) on the inner face of the membrane by causing an exchange of a GTP for G DP at a nucleotide binding site (Dohlman et al., 1991; Hollenberg, 1991; Simon et al., 1991). This activated G protein then may directly cause alteration of an ion channel in the membrane (Neer and Ciapham, 1988). activate or inactivate adenyl cyclase (Freissmuth et al., 1989) and thus alter the levels of cAMP in the cell, or activate a phospholipase in the cell membrane leading to the production of second messengers from the membrane phospholipids (Ferguson and Hanley, 1991). The latter include diacylglycerol (DAG) and inositol triphosphate (IP3) produced from phosphatidylinositol 4,5-bisphosphate (PIP2) (Berridge and Irvine, 1989), and arachidonic acid or D AG or phosphatidic acid from phosphatidyl choline (Exton, 1990). The second messenger cAMP activates protein kinase A (McKnight. 1991), and D A G activates protein kinase C (Bell and Burns, 1991), both of which have multiple forms that catalyse phosphorylation of various cellular proteins. This phosphorylation may either activate or inactivate an enzyme such as activation of glycogen phosphorylase (Sutherland, 1972), open or close an ion channel, or alter transcription factors. In the latter case. protein kinase A is known to phosphorylate a cyclic-AMP-response-element-binding protein (CREB) that then binds an'd activates certain CAMP-inducible genes (Habener, 1990; Karin, 1991). Protein kinase C regulates both phosphorylation and dephosphorylation of the AP-1 transcription factor complex (which includes c-fos and c-jun), thus having both positive and negative effects on gene transcription. These factors may also form heterodimers with steroid hormone receptors (Karin, 1991; Schiile and Evans, 1991; Miner and Yamamoto, 1991). Inositol triphosphate acts on the endoplasmic reticulum to cause the release of calcium (Berridge and Irvine, 1989). Calcium then combines with the ubiquitous protein calmodulin to modulate various cell activities including the activation of a specific calcium-calmodulin protein kinase which regulates many different cellular functions (Hanson and Schulman, 1992). Besides the G-protein coupled receptors, there are transmembrane receptors for peptide hormones that, when ligand is bound, change conformation so that a catalytic site on the cytoplasmic face of the receptor molecule is activated and thus generates a second messenger. These include the insulin and growth factor receptor tyrosine kinases (Yarden and Ullrich. 1988) and the receptor guanylyl cyclases which generate cGMP (Chinkers and Garbers, 1991). The tyrosine kinases cause both tyrosine autophosphorylation and thus inactivation of the receptor and tyrosine phosphoryla-

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tion of other intracellular proteins. cGMP may also be produced by activation of a cytoplasmic guanylate cyclase through the action of arachidonic acid or nitric oxide (Chinkers and Garbers, 1991; Goy, 1991) or decreased by a G-protein-mediated activation of cGMP phosphodiesterase as in the photoreceptors of the eye (Stryer, 1991). 2 Embryonic actions of juvenile hormone As discussed above, J H normally is present during the last third of embryonic development due to secretion by the newly formed corpora allata (Briining et al., 1985; Biirgin and Lanzrein, 1988; Cusson et al., 1991). It appears to be important for both the first instar larval cuticle production and for the normal development of the gut (Dorn, 1982, 1990; Bruning and Lanzrein, 1987). Whether organogenesis of other tissues is also affected is not known. Also, the type of action that J H has in promoting this development is not known. In Locusta J H is produced earlier in development by the serosa which also has a JH-binding protein (Hartmann et al., 1987), but its role at this time has not been determined. J H esterase activity is high at the outset of embryonic development in both the cricket, Achaeta domesticus (Roe et al., 1987), and in Manduca (Share et al., 1988), then falls to low levels. This rids the egg of maternal J H and allows normal development since J H can adversely affect embryonic development. When given in high doses to either the female during oogenesis or to the freshly oviposited eggs, J H and J H analogs (JHAs) prevent normal development (Riddiford, 1972; Staal, 1975; Kelly and Huebner, 1986). After treatment of the female, embryogenesis may be blocked at sometime shortly after the blastoderm stage when the germ band begins development (reviewed in Riddiford, 1972). In the linden bug, Pyrrhocoris apterus, the hormone is most effective if given to the female during the postvitellogenic, prechorionic stage of egg maturation, but the mode of action is unknown. By contrast, treatment of the eggs of many species just after oviposition with J H results in embryos that show defects in dorsal closure and the movements associated with blastokinesis (Riddiford, 1972; Wall, 1974; Enslee and Riddiford, 1977; Injeyan et al., 1979; Kelley and Huebner, 1986). In Pyrrhocoris, failure of blastokinesis appears to be due to lack of formation of a band of apical microfilaments in some cells of the serosa (Enslee, 1975). A complete band is necessary for the contraction of this sheet which helps to pull the embryo through blastokinesis (Enslee and Riddiford, 1981). Also, various aspects of organogenesis such as the shortening of the nerve cord in Pyrrhocoris may be disrupted (Riddiford, 1972). Again, the mode of action of J H in creating these abnormalities is unknown. Specific protein differences have been seen in treated Rhodnius eggs (Kelly and Huebner, 1987), but how these relate to the developmental effects is unclear.


3

3.1

225

Premetamorphic actions of juvenile hormone REGULATION OF CELLULAR COMMITMENT BY J U V E N I L E HORMONE

The primary role of J H during larval life is to prevent metamorphosis. This action is manifest at every moult, and J H must be present at the outset of the ecdysteroid rise for the moult in order for the cells to retain their larval characteristics. During the final larval instar when JH is low or absent, changes occur that commit the cell to metamorphosis so that even if later exposed to ecdysteroid in the presence of JH, it can no longer respond with a larval moult. These changes d o not occur at the same time in all tissues, in all cells within a tissue, or for all activities of a single cell so that application of exogenous J H during this critical period results in mosaic animals with characteristics of two different stages (Wigglesworth, 1940; Williams, 1961; Willis et al., 1982; Nijhout, 1983; Ohtaki el al., 1986; Kremen and Nijhout, 1989). In Lepidoptera, the imaginal discs first become committed, then the epidermis that is to assume a new fate, then the larval tissues that die at metamorphosis (reviewed in Kremen and Nijhout, 1989). Within the epidermis the cells become committed according to their position in the animal and within the segment (Truman et al., 1974; Kremen, 1989), and within cells commitment for the surface patterning and for pigmentation occurs at different times (Willis et a l . , 1982). Also, the viscera such as the gut, the muscles, silk glands and the nervous system become committed in a particular pattern (Riddiford, 1972; Sehnal and Schneiderman, 1973; Cymborowski and Sehnal, 1980; Ohtaki et al., 1986; Weeks and Truman, 1986a,b; Sehnal and Akai, 1990). These changes in commitment are often, but not always, correlated with small surges of ecdysteroid in the absence of JH as will be outlined in the sections below. Most studies have not tested the necessity of this ecdysteroid for the change. In vitro studies with Manduca abdominal epidermis clearly demonstrated that low levels of 20-hydroxyecdysone (20E) were essential for the change to pupal commitment in that tissue (Riddiford, 1976). Also, ecdysteroid infusions into isolated abdomens have shown that the hormone is necessary for the changes in commitment of the muscles and nervous system of Manduca (Weeks and Truman, 1986b). By an extensive series of cautery experiments, Kremen (1989) demonstrated that the change of commitment in the epidermis of the butterfly, Precis coenia, begins in certain foci in each segment and requires intercellular communication. In this species there are small, but statistically nonsignificant, fluctuations in ecdysteroid titre during the final intermoult period when changes in commitment are occurring (Kremen and Nijhout, 1989). Therefore, these authors hypothesize that the decline of JH allows the onset of intercellular communication that leads to pupal commitment. Exogenous JH then is thought to delay or interrupt this process and thus to prevent the

L. M. RlDDlFORD

226

change. Since exogenous 20E that caused an early moult had no effect on the sequential pattern of pupal commitment, they suggest that the decline of JH may be sufficient to cause the change of commitment. The critical test of this hypothesis is to determine whether larval epidermis isolated from hormonal sources becomes committed to metamorphosis so that it will form pupal cuticle when exposed to a moulting concentration of ecdysteroid in the presence of JH. This has not been done with Precis epidermis. The following sections will deal with the individual larval tissues where we are beginning to learn about the cellular and molecular basis of this action of JH in guiding cellular commitment. The focus will be on those systems in which specific proteins and/or genes have been studied. Juvenile hormone is critical to the regulation of the stage specificity of both genes expressed during larval life and those that are first expressed at metamorphosis. During larval life, its presence during the moult allows the surge of ecdysteroids to suppress ongoing expression of intermoult genes but prevents their permanent repression. Consequently, when ecdysteroids disappear, these genes can be expressed again. At metamorphosis ecdysteroids act in the absence of JH, and larval-specific genes are permanently suppressed. Also, at this time new genes specific to the pupa or adult are committed to later expression by low levels of ecdysteroids acting in the absence of JH.

3.2

EPIDERMIS

The epidermis makes the cuticle that lies above it and is responsible for pigmentation. The structure of this cuticle is dependent on the proteins deposited and their interaction with chitin. Different proteins are needed when different cuticular structures are to be formed, whether it be during the moult or during the intermoult period (Doctor et al., 1985; Wolfgang and Riddiford, 1986; Willis, 1991; Apple and Fristrom, 1991). Moreover, stage-specific cuticles are a mosaic of both conserved and new stage-specific proteins as first clearly defined in the wild silkmoth, Hyalophora cecropia (Cox and Willis, 1985, 1987). 3.2.1 Regulation of larval cuticle gene expression in Manduca One of the best systems for the study of the cellular and molecular basis of the morphogenetic action of JH has been the epidermis of the tobacco hornworm. This epidermis responds to hormonal manipulations in vitro as in vivo. When larval epidermis is exposed to 20E in the presence of JH, it forms a new larval cuticle after the ecdysteroid is removed; when exposed in the absence of JH, it will first become pupally committed, then produce a pupal cuticle (Truman et al., 1974; Fain and Riddiford, 1976; Mitsui and


227

Riddiford, 1978; Riddiford, 1976, 1978; Hiruma et al., 1991). Moreover, several of the larval cuticle genes have been isolated and their hormonal regulation studied. Since these studies are contributing to a deeper understanding of our knowledge of the molecular basis of JH action, they will be briefly summarized below. 3.2.1.1 LCPl4 The LCP14 gene encodes a 14 kDa endocuticular protein and is expressed during every larval growth phase, but not during the moult (Rebers and Riddiford, 1988; Fig. 4). When the epidermis becomes pupally committed in response to low ecdysteroid in the absence of JH on the final day of feeding (Riddiford, 1978), LCP14 mRNA disappears and never reappears during or after metamorphosis. The LCP14 mRNA present in 4th instar epidermis rapidly disappeared during exposure to 20E in vitro (Hiruma et a / . , 1991). It then reappeared some time after the 20E was removed if J H had been present in the cells at the time of exposure to 20E. This suppressive action of 20E required protein synthesis, likely of an ecdysteroid-induced transcription factor(s).

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Thus, the ultimate fate of this larval-specific gene that is expressed during the intermoult periods depends on the presence or absence of J H at the time that 20E suppresses its expression. When J H is present, the gene can be reexpressed once the ecdysteroid-induced suppressive protein(s) disappear. When J H is absent, the gene somehow becomes permanently inactivated by ecdysteroid. Whether J H acts directly on the gene itself, modulates the type of EcR and/or ecdysteroid-induced transcription factors present, or modifies chromatin structure is not known (see further discussion below).

3.2.1.2 LCP 14.6 The gene encoding the 14.6 kDa larval endocuticular protein has a different and intriguing developmental expression pattern (Riddiford et al., 1986; Fig. 4). The mRNA was found throughout the segment in the larva with two peaks of expression-just before ecdysis and during the intermoult. After pupal commitment, expression was restricted to the flexible intersegmental regions and occurred only just before pupal and adult ecdysis. In situ analysis (Riddiford, 1991; Riddiford and Poon, unpublished) shows that the mRNA is present in all larval epidermal cells except the hair cells, but in the pharate pupa is present only in those cells making the intersegmental cuticle. In the pharate adult, the mRNA is highest in cells around the muscle attachment sites on the flexible cuticle. Thus, through metamorphosis as the cells respond to ecdysteroid in the absence of JH, expression of this gene becomes progressively spatially and temporally restricted. Just like LCPl4, this gene was suppressed by 20E during a larval moult or at pupal commitment in vitro (Riddiford, 1986; Riddiford et al., 1986). But unlike LCP14, expression of LCP14.6 in 5th instar epidermis could also be suppressed by the J H A methoprene in the absence of ecdysteroid. Whether this suppression depends on JH-induced protein synthesis has not been tested. In vivo levels of LCP14.6 mRNA are highest when JH is low, but the role of J H in its intermoult regulation is unclear. 3.2.1.3 LCP16/17 On the last day of feeding in the final (5th) larval instar, Manduca epidermis synthesizes a new set of cuticular proteins that are correlated with a decreased spacing of the cuticular lamellae (Wolfgang and Riddiford, 1986) and an increase in the flexural stiffness of the cuticle (Wolfgang and Riddiford, 1987). At least three of these new proteins (16, 16.3 and 17 kDa) are encoded by the multigene family LCP16/17 (Horodyski and Riddiford, 1989). The LCP 16/17 genes have a narrow temporal patttern of expression as the mRNAs are expressed only on the final day of feeding in the last instar (Horodyski and Riddiford, 1989; Fig. 4). The onset of this expression was found to be dependent on the small rise of ecdysteroid that occurs on day 2 (Wolfgang and Riddiford, 1986) and was prevented by exogenous JH. Induction of the mRNA by 20E was prevented by inhibition of protein


229

synthesis (Riddiford, 1991), indicating that a new ecdysteroid-induced transcription factor(s) may be necessary. Whether J H prevents the appearance of this factor, interferes with some ecdysteroid-induced change in chromatin structure that enables the activation of previously unexpressed genes, or has some suppressive effect directly on these genes is unknown. 3.2.2 Regulation of larval and pupal cuticle gene expression in higher Diptera In the higher Diptera, the 3rd instar larval cuticle transforms into a hardened puparial case at the onset of metamorphosis that protects the developing pupa. The larval epidermal cells of the head and thorax then die while those of the abdomen make the pupal cuticle (Bhaskaran, 1972; Madhavan and Schneiderman, 1977). In DrosophiZa the urea-extractable cuticular proteins of the 1st and 2nd instar larvae are the same, but a new set is made in the 3rd (final) instar (Chihara et al., 1982) with two of the major proteins appearing earlier than the others (Kimbrell et al., 1988, 1989). Presumably these new proteins are the type that can crosslink later with the various sclerotizing agents formed in response to the pupariation peak of ecdysteroids (Hopkins and Krarner, 1992). Interestingly, the water-soluble proteins of the cuticles of penultimate and final larval instar Sarcophaga bullata larvae appear to be the same (Willis, 1991), indicating that only some proteins change. The hormonal control of this switch in cuticular protein synthesis is unclear. In Drosophila the switch is correlated with a decrease in the JH titre (Bownes and Rembold, 1987; Sliter et af., 1987) and an increase in J H metabolism in the early 3rd instar (Campbell et al., 1992). However, continuous exposure to J H analogues in the diet had little effect on puparium formation (Riddiford and Ashburner, 1991), but whether these new cuticular mRNAs appeared under these conditions has not been determined. 3.2.3 Regulation of pupal and adult cuticle genes Studies of translatable mRNAs from the mesonotum of the waxmoth Galleria mellonella have revealed a changing pattern of expression which is modulated by JH (see Wolbert and Schafer, 1991 for a review). Immediately after pupal ecdysis, mRNAs for pupal endocuticular proteins are present, then within a day or so they disappear. This disappearance coincides with the loss of sensitivity to exogenous JH to cause formation of a second pupal cuticle. Exogenous J H delays but does not prevent the disappearance of the pupal messengers. During the critical period of JH sensitivity in addition to changes in cuticular proteins, synthetic patterns of both cytoplasmic and nuclear

230

L. M. RlDDlFORD

proteins change (Wolbert and Schafer, 1991). Yet J H seems to have little effect on these changes. At least one cytoplasmic and one nuclear protein that may be associated with commitment to adult differentiation (based on the timing of their appearance) do not appear in the presence of JH. The function of these proteins is not known. Whether one should expect to see electrophoretic differences at this adult commitment stage is unclear since hormone receptors and transcription factors are minor constituents of the cellular proteins and therefore may not be detectable without specific probes. Later coincident with the rise of ecdysteroid and the production of the new cuticle, new mRNAs appear and many later disappear in a particular temporal sequence, presumably corresponding to the type of cuticle being made (whether epi-, exo-, or endocuticle). In the wild silkmoth, Antheraea polyphemus (Sridhara, 1985), and in the beetle, Tenebrio molitor (Bouhin et al., 1992), these new mRNAs are not observed when J H is given just after pupal ecdysis before the onset of adult development. In Tenebrio, one of these new mRNAs encodes an adult-specific cuticular gene. Instead, the mRNAs are similar to those found in epidermis making pupal cuticle as would be expected since a second pupal cuticle is formed under these conditions. 3.2.4 Regulation of larval pigmentation

In many insects larval pigmentation is under the control of J H and is often associated with environmental signals such as population density (see Nijhout and Wheeler, 1982; Riddiford, 1985; Pener, 1991 for reviews). The critical period of J H sensitivity is also during the ecdysteroid rise for the larval moult, but later than that for the determination of the type of moult. The cellular and molecular basis of this action of J H is best known in Manduca and will be summarized below. 3.2.4.1 Insecticyanin Manduca larvae have a blue biliprotein, insecticyanin, in the epidermis and haemolymph (Cherbas, 1973; Riley et al., 1984; Trost and Goodman, 1986; Goodman et al., 1987). The larval epidermis during the intermoults synthesizes, stores, and secretes two major forms of insecticyanin, INS-a (PI 5.5) and INS-b (PI 5.7) (Kiely and Riddiford, 1985; Riddiford et al., 1990) which are encoded by different genes (Li and Riddiford, 1992). Only INS-b is present in the haemolymph. During larval life insecticyanin mRNA in the epidermis is high during the intermoults, then disappears during the moults due to the action of 20E (Riddiford et al., 1990) just as seen for LCP14 (Fig. 4). On the final day of feeding in the last instar, synthesis of INS-a ceases, whereas that of INS-b continues until wandering (Kiely and Riddiford, 1985; Riddiford et al., 1990) (Fig. 4). Exposure of day 1 5th instar larval epidermis to 20E in vitro in two


23 1

pulses as normally seen in vivo (Wolfgang and Riddiford, 1986) caused the differential disappearance of INS-a mRNA (Li, Hiruma and Riddiford, unpublished). Presumably differences in the promoter regions of the two genes are responsible for this differing sensitivity to ecdysteroid in the absence of JH. J H during a larval moult not only determines whether or not insecticyanin mRNA will reappear after the moult, but also the level of its accumulation in the epidermis during the 5th instar (Goodman et al., 1987). The critical period for this quantitative effect of JH is at the time of head capsule slippage near the peak of the ecdysteroid rise (Fig. 4). When JH is absent at this time, less insecticyanin accumulates in 5th instar epidermis; JH applications restore normal levels (Goodman et al., 1987). Preliminary experiments suggest that the decreased accumulation is due to the delayed resumption of insecticyanin mRNA synthesis after ecdysis in those larvae lacking JH during the critical period (Li and Riddiford, unpublished). The molecular basis for this delay has not yet been determined. 3.2.4.2 Cuticular melanization Normally, Manduca larvae have a transparent larval cuticle except for a set of black markings on the dorsal abdomen. During the larval moult, the absence of J H at the time of head capsule slippage causes the later melanization of the new cuticle just before ecdysis (Truman et al., 1973; reviewed by Riddiford and Hiruma, 1988). The melanin is formed from dopamine in granules deposited into the new endocuticle which contain a specific prophenoloxidase and other enzymes necessary for converting dopamine. to melanin (Hiruma and Riddiford, 1984, 1988; Hiruma et al., 1985; Hopkins and Kramer, 1991). Then as the ecdysteroid titre declines, the phenoloxidase is activated and the dopamine formed by the action of dopa decarboxylase (DDC) on dopa is converted to melanin. 3.2.4.2.1 Granular phenoloxidase. The granular phenoloxidase (PO) responsible for melanization is immunologically distinct from the wound and haemolymph phenoloxidases (Hiruma and Riddiford, 1988). When JH is absent at the time of head capsule slippage, PO synthesis begins 6 h later in the epidermis as the ecdysteroid titre falls. The proenzyme is deposited along with other granular proteins into the newly forming cuticle over about the next 12-14 h. Activation of the enzyme then occurs so that melanin appears about 3 h before ecdysis. Juvenile hormone prevents the onset of phenoloxidase synthesis if given at head capsule slippage which occurs near the peak of the ecdysteroid titre (Fig. 4). Once synthesis has begun, the addition of J H prevents any further increase in rate of synthesis (Hiruma and Riddiford, 1988). Thus, J H either directly or indirectly suppresses the later onset of synthesis of this unique phenoloxidase and the associated proteins of the granules in response to the moulting surge of ecdysteroids. This action then is typical of JH in

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preventing the ecdysteroid-induced expression of new genes. By contrast, the effect of J H on the increasing rate of ongoing synthesis may be novel. In the larval lepidopteran fat body, J H suppresses the appearance of mRNAs for certain storage proteins, but has no effect on the level once it appears (see Section 3.3.3 below). Further clarification requires isolation of the phenoloxidase gene in order to determine whether JH is modulating transcription or translation in its effect on ongoing synthesis of the enzyme. 3.2.4.2.2 Dopa decarboxylase. Dopa decarboxylase (DDC) is essential for both melanization and sclerotization of the cuticle so activity is normally high at the end of the moult (Hopkins and Kramer, 1992). To provide the extra dopamine required for melanization, D D C activity is two-fold higher in epidermis of Manduca larvae that are destined to melanize (Hiruma and Riddiford, 1985). In Manduca, DDC mRNA appears during the latter part of the moult, peaking about 10 h before larval ecdysis and then again just before pupal ecdysis (Hiruma and Riddiford, 1990; Pedersen, Hiruma, and Riddiford, unpublished; Fig. 4). Maximal expression was two-fold higher in melanizing allatectomized larvae and could be returned to normal levels by application of JH I to these larvae at head capsule slippage (Hiruma and Riddiford, 1990). Thus, JH at the time of head capsule slippage during the moult quantitatively regulates the later level of expression. The regulation of DDC expression during the moult is complex. In vitro studies with 4th instar epidermis have shown that 20E is necessary for later induction of D D C mRNA transcription, but its presence suppresses that transcription so that no DDC mRNA is seen until the ecdysteroid titre declines (Hiruma and Riddiford, 1990). A simple hypothesis to account for this dual action of 20E is that the hormone induces two transcription factors with differing half-lives (Hiruma and Riddiford, 1993; Fig. 5 ) . Factor A has a long half-life and is necessary for induction of D D C mRNA synthesis, but the presence of the second factor, B, prevents the expression of DDC. When the ecdysteroid titre declines, B disappears first due to its shorter halflife. Then DDC can be expressed until factor A also disappears. Consequently, one obtains a pulse of DDC expression. J H is hypothesized to have its quantitative effect on DDC levels by influencing either the transcription or translation of factor A . Experiments have shown that the suppression of D D C by 20E is due to a short-lived ecdysteroid-induced protein(s) (Hiruma and Riddiford, 1993 and unpublished). At the time of head capsule slippage near the peak of the ecdysteroid titre, no DDC mRNA is present. When cycloheximide, a protein synthesis inhibitor, was given at this time either in vivo or in vitro, high levels of DDC mRNA were observed 4 h later. Extracts of epidermal nuclei at the time of head capsule slippage but not at the time of maximal DDC expression contain a factor that binds to a single region in the 5' promoter of the DDC gene. Presumably then the former extract contains


Ec + EcR

233

+Ec-EcR

+- -

DDC

f-

Ec-EcR FIG. 5 Hypothetical scheme for the action of 20-hydroxyecdysone (Ec) and juvenile hormone (JH) in the regulation of dopa decarboxylase (DDC) expression during a larval moult (from Hiruma and Riddiford, 1993). A and B are transcription factors. R, receptor. See text for detajls.

the ecdysteroid-induced negative regulatory factor(s) B, and studies are under way to determine its nature. Several other fragments of the promoter region bind extracts from both stages, indicating that there may be more than one binding site and/or ecdysteroid-induced positive regulatory factor necessary for activation of the DDC gene. Once more is known about these factor(s), the influence of J H on their level can be determined. 3.2.5

Imaginal discs and other imaginal precursors

During larval life imaginal discs of Lepidoptera and Diptera proliferate but form no cuticle. This proliferation occurs continuously, irrespective of the moult cycle (Bryant and Simpson, 1984). Proliferation of transplanted larval Drosophila discs also occurs in adult females fed a protein diet, but not in starved females nor in males (Hadorn and Garcia-Bellido, 1964; GarciaBellido, 1964; Schubiger, 1973). Even in this situation, the hormonal conditions that promote proliferation are unclear. Females have higher ecdysteroid levels than males (Schwartz et al., 1989) but the level in females after 2 days of starvation is unaltered (Bownes, 1989). Also, the JH levels

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after 2 days of starvation are similar (Bownes, 1989). In no case is the haemolymph level of either ecdysteroids or J H known and whether the hormone levels remain normal after further starvation has not been studied. The prolonged effects may be important since starved Manduca larvae show first an increase in J H titre in the haemolymph, followed in 3-4 days by a rapid decline (Cymborowski et al., 1982). Proliferation has been difficult to achieve in vitro, but will occur in wounded whole discs if the medium contains methoprene and insulin (Davis and Shearn, 1977). With late larval or prepupal discs, insulin appears to enhance ecdysteroid action on evagination but to inhibit its stimulation of pupal differentiation (Martin and Shearn, 1980). No further studies have been done with methoprene or with a natural J H to determine its necessity in this system. At metamorphosis the imaginal discs begin to differentiate into the various pupal precursors of the adult antennae, eyes, legs, wings and genitalia. In the Lepidoptera the discs become committed to pupal differentiation very early in the final instar (reviewed in Kremen and Nijhout, 1989) long before the body epidermis. Thus, J H application to final instar lepidopteran larvae can cause the formation of supernumerary larvae with pupal wing discs. By contrast, J H has no such effect when given to either early or late stage Drosophila larvae (Postlethwait, 1974; Riddiford and Ashburner, 1991) although at high concentrations in vifro, it prevents the pupal response of discs to 20E (Chihara et al., 1972; Chihara and Fristrom, 1973; Doctor and Fristrom, 1985). Indeed, the only effect of raising larvae on diet containing a high concentration of a persistent J H A . such as pyriproxyfen was to prevent adult development (Riddiford and Ashburner, 1991). The discs evaginated and formed a pupal cuticle, but adult differentiation was prevented. By contrast, exposure to the same concentration beginning in the mid-2nd instar when the discs are becoming competent to metamorphose had no effect on adult differentiation of imaginal disc derivatives. Thus, the effect was not due to the persistence of the JHA to the time of the onset of adult differentiation. Rather the timing of the effect was correlated with the onset of proliferation of the imaginal discs in the mid- to late 1st instar (Madhavan and Schneiderman, 1977) at the time of the ecdysteroid rise for the moult to the second stage (Kraminsky et al., 1980). The presence of very high J H during this early proliferative phase had little effect on the acquisition of competence to begin metamorphosis but prevented the later ability to respond to ecdysteroid by adult differentiation. Juvenile hormone also inhibits adult differentiation of the abdominal histoblasts in Drosophila (Ashburner, 1970; Madhavan, 1973; Postlethwait, 1974) and Sarcophaga bullata (Bhaskaran, 1972) when applied between pupariation up to shortly after pupation as signalled by head eversion. This is the time of histoblast proliferation followed by spreading over the


235

abdomen in preparation for later production of the adult cuticle (Madhavan and Schneiderman, 1977; Roseland and Schneiderman, 1979). To be most effective, J H must be given during the proliferative phase (Postlethwait, 1974). In both Sarcophaga (Bhaskaran, 1972) and Drosophilu (Currie and Riddiford, unpublished), the exogenous J H had no effect on the proliferation of the histoblasts, but in some cases prevented degeneration of the larval cells and the spreading of the histoblasts as well as the later differentiation. Thus, the presence of J H may be permissive for imaginal cell proliferation in the larva. However, excess levels at some time during this proliferation can have profound effects much later at the time of the onset of adult differentiation. Whether this action of JH is dependent on the action of ecdysteroid as are its other morphogenetic actions or whether it simply inhibits some cellular event that is necessary for their later response to ecdysteroid during the adult moult are important questions.

3.3

FATBODY

The fat body serves to maintain homeostatic levels of nutrients in the haemolymph and to synthesize various haemolymph proteins as well as vitellogenins in the adult. During the larval feeding stages, most of the incoming nutrients are utilized for growth. Then during the final larval instar of both hemimetabolous and holometabolous insects, the fat body synthesizes and secretes high amounts of several large, usually hexameric proteins often referred to as storage proteins (see Levenbook, 1985; Kanost et al., 1990; and Telfer and Kunkel, 1991 for reviews). In holometabolous insects, these proteins are stored in the pupal fat body in the form of large crystalline granules for later use during the pupal-adult transformation (Dean et al., 1985). Studies are just beginning on the hormonal regulation of these haemolymph proteins. Only those specifically regulated by J H will be considered here. 3.3.1 Arylphorin The arylphorins are a class of storage proteins rich in aromatic residues (tyrosine, phenylalanine) (Telfer et al., 1983) and are used in the formation of new cuticle of various stages (Konig et al., 1986; Webb and Riddiford, 1988a; Peter and Scheller, 1991) as well as for the formation of various adult structures such as flight muscle (Levenbook and Bauer, 1984). In higher Diptera, arylphorin is only synthesized by the fat body of final instar larvae (Kanost et al., 1990), and the remnants of that fat body in the adult (Benes et al., 1990). The role of JH in suppressing expression in the earlier larval stages has not been studied.

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In the Lepidoptera, arylphorin mRNA is present and the protein is synthesized during the feeding phase of the larval instars, but not during the moults (Webb and Riddiford, 1988a, b; Memmel et al., 1988; Fujii et al., 1989; Fig. 6). Just as with the larval cuticle proteins LCP14 and LCP14.6 (see Section 3.2.2), the high ecdysteroid titre during the moult causes this disappearance of arylphorin mRNA (Webb and Riddiford, 1988b). The reappearance of the mRNA and resumption of synthesis occurs at some time after each larval ecdysis and was shown to be dependent on incoming nutrients (Memmel et al., 1988; Webb and Riddiford, 1988a,b). Presumably this resumption is dependent on the presence of JH at the time of the moult since arylphorin mRNAs disappear early in metamorphosis during or just after the pupal moult, depending on the species. In Bombyx an mRNA for a small 30 kDa larval-specific haemolymph protein has a similar developmental expression pattern to that of arylphorin through larval life-high during the intermoults, suppressed during the

ECDYSIS

r o l l

' 2 ' 3 ' 4 1 0 '

1-4Ih-~-Sth-

d&' ' '

4 '

' ' '

Met rich

SP I

' '

15' ' ' 'IS' ' ' IS' ' ' ' Irn' ' ' ' ' -davaloplnp 'adull -I-adult

@ - .@ & Qp& < 0

@ID

Arylphorin

a'o'

I-pharata -I-pupa pupa a.

4 , -3(I

p

.@

'L

-

30K

FIG. 6 Expression of genes in the fat body of Lepidoptera during larval development and metamorphosis. The titre is that of Manduca sexta (Fig. 2b); the schematic of fat body development is based on Calpodes efhlius (Dean et al., 1985); arylphorin and methionine (met)-rich mRNAs are from Manduca sexta (Webb and Riddiford, 198%); SP1 (storage protein 1) (Tojo et al., 1981; Sakurai ef al., 1988), BmLSP (larval-specific protein) (Fujiwara and Yamashita, 1991), and 30 K (Sakai et al., 1988; Mori et al., 1991) from Bombyx mori.


237

moults (Fujiwara and Yamashita, 1990, 1991). mRNA expression declines early during the final instar at which time other JH-suppressible 30 kDa proteins appear (see Section 3.3.3 below). Whether the disappearance of JH in the final larval instar is responsible for this decline has not been studied. 3.3.2 Methionine-rich storage proteins The methionine-rich storage proteins are found primarily in the Lepidoptera and often show sex specificity (Kanost et al., 1990). In Bombyx, penultimate instar larvae of both sexes synthesize the methionine-rich SP1 at low levels; then in the final instar only females do so and at much higher levels (Tojo et al., 1981; Mine et al., 1983; Sakurai et al., 1988; Fig. 6). In sexual mosaics the cessation of SP1 synthesis occurs only in the male cells (Mine et al., 1983), indicating cell autonomous, sex-specific control. The sex-specific changes can be prevented by exposure to exogenous J H (methoprene) early in the penultimate instar, but not after the critical period for J H in the final larval moult (Kajiura and Yamashita, 1989). Also, before the final larval moult, methoprene suppressed SP1 synthesis in both sexes; whereas in the final instar it promoted SP1 synthesis in males, but had no effect in females. Thus, the hormonal milieu of this final larval moult interacts somehow with sex-specific factors to commit the fat body to enter its preparative phase for metamorphosis. In Manduca female larvae, the mRNAs for the methionine-rich proteins and the proteins themselves are not found until day 2 of the final instar; in males synthesis begins later at the .onset of wandering (Ryan et a f . , 1985; Webb and Riddiford, 1988a; Corpuz et al., 1991) (Fig. 6a). The decline of J H is necessary for this appearance (Riddiford and Hice, 1985; Webb and Riddiford, 1988b; Corpuz et al., 1991). Transcripts for two different proteins initially were suppressed after treatment with methoprene, then with time as the JH declines, SPlA but not SPlB reappeared (Corpuz et a f . , 1991). No detailed study of this interesting phenomenon has been done. 3.3.3 Other JH-suppressible storage proteins In various Lepidoptera ( B o m b y x (Plantevin et al., 1987; Bosquet et al, 1989a, b), Trichoplusia ni (Jones et al., 1988, 1990) and Galleria (Kumaran et al., 1987; Memmel and Kumaran, 1988; Memmel et al., 1988)) and in final instar larvae of the Colorado potato beetle, Leptinotarsa decemlineata (Koopmanschap et al., 1992), new serum proteins appear in the mid-feeding stage of the final larval instar. In Bombyx these are 30 kDa proteins encoded by several genes which show similar, but individual, patterns of expression (Sakai et al., 1988; Bosquet et al., 1989a,b; Mori et al., 1991). The mRNAs for these proteins normally appear when the J H titre falls, and J H application during or just after the moult to the final instar prevents

238

L.

M. RlDDlFORD

(Trichoplusia, Galleria and Leptinotarsa) or retards ( B o m b y x ) their appearance. In vivo and in vitro experiments suggest that this suppressive effect of J H is directly on the fat body (Ray et al., 1987; Bosquet et al., 1989a). However, in Galleria (Ray et al., 1987), once these RNAs appear, JH is no longer suppressive; further transcription then is only suppressed by ecdysteroid. Thus, all these JH-suppressible proteins and the methioninerich protein in Manduca are like the LCP16/17 cuticular proteins in that they are indications of impending metamorphosis. Whether ecdysteroids are necessary to activate these genes in the fat body after the JH titre declines as they are for LCP 16/17 expression has not been tested. 3.3.4 JH-inducible larval proteins The haemolymph of midge larvae contains haemoglobins to improve their respiratory capacity in their low oxygen environment. In Chironomus thummi, there are multiple electrophoretic forms of these haemoglobins (Vafopoulou-Mandalos and Laufer, 1982) encoded by at least five genes (Antoine et al., 1987). These are synthesized by the fat body during the intermoult periods, but not during the moults, and then synthesis ceases at metamorphosis (Schin et al., 1979) (Fig. 6 ) . Exposure of fat body to 20E in vitro showed that this hormone suppressed synthesis of all but one form of haemoglobin (Vafopoulou-Mandalos and Laufer, 1984). Whether this suppression is entirely at the level of transcription of the specific RNAs as is arylphorin is unclear (see Section 3.3.1; Fig. 6). Superimposed on this general pattern is a changing developmental pattern of the various forms synthesized (Vafopoulou-Mandalos and Laufer, 1982, 1984). Of particular interest are two haemoglobins that normally do not appear until the 4th day of the final larval instar. Synthesis of these two was induced precociously in the final instar by J H (methoprene) both in vivo and in vitro and was due to an increase in translatable mRNA (VafopoulouMandalos and Laufer, 1984). In vitro the induction was weak as compared to the induction seen in vivo, but curiously occurred only with body walls (integument, muscle, and fat body) of day 0 final instar larvae and not with body walls from either penultimate instar or mid-final instar larvae. Whether these differences are true developmental changes or are due to some artefact of the culture system is not certain. Since these two haemoglobins normally do not appear until mid-final instar, it seems paradoxical that J H should activate their transcription. Now that the haemoglobin genes are available (Antoine et al., 1987), this potentially interesting effect of J H should be studied further. 3.3.5 Larval serum proteins in the Hemimetabola In the cockroach Blattella germanica (Duhamel and Kunkel, 1987) and in


239

Locusta (Wyatt et al., 1992a), there are also hexameric haemolymph proteins that cycle with the moulting cycle. Whether they are controlled at the level of synthesis or uptake is unclear. In the bean bug, Riptortus clavatus, the two subunits of the hexameric cyanoprotein are synthesized only during the intermoult periods with each subunit showing its own pattern of synthesis (Chinzei et al., 1992). These cycling patterns in various hemimetabolous insects are reminiscent of that of arylphorin, so their hormonal regulation is likely similar. The difference between the Hemimetabola and the Holometabola in regulation of these hexameric proteins is seen at metamorphosis. At this time in the Hemimetabola, there may not be a permanent cessation of synthesis, but rather an alteration in the pattern of synthesis. For instance, in Locusta larval-specific storage protein 1 (LSP1) and persistent storage protein (PSP) are present in the same proportion in each of the final three larval instars (Wyatt et al., 1992a). Late in the final instar LSPl increases relative to PSP, apparently due to differing sensitivities of the two genes to the lack of JH. Application of the J H A pyriproxyfen suppressed the increase of LSP, but had little effect on PSP. In the adult, levels of LSPl decline rapidly. By contrast, PSP in the adult remains high due to continued synthesis; at this time, synthesis is stimulated by J H (Wyatt et al., 1992b). Similarly, in Riptortus, both subunits of the cyanoprotein are made in the nymphal instars by both sexes. Then in the adult reproducing female only one is made, while in the adult male, neither is synthesized (Chinzei er al., 1992). Thus, the metamorphic moult in the absence of JH does not permanently inactivate these genes, but changes the mode of their regulation. One possible difference may be the type of transcription factors available or the possible appearance of a new type of J H receptor.

3.4

MUSCLE

At metamorphosis larval muscles either die or are restructured and/or reoriented to acquire a new role in the adult. For instance, certain abdominal intersegmental muscles of Lepidoptera persist unchanged through adult development to be used during eclosion, then die thereafter (Lockshin, 1985). New muscles specific for adults such as the flight muscles appear at metamorphosis. All of these events are under the control of JH, but nothing has been done at the cellular or molecular level to define its mode of action. Certain key findings will be outlined below. The abdominal prolegs that are characteristic of lepidopteran caterpillars are larval-specific. In Manduca the epidermis which forms the crochets (hooks) on the tips of these prolegs becomes committed to die early in the final larval instar before the remainder of the epidermis and shows signs of cell death by the onset of wandering (Fain and Riddiford, 1977). The

L.

240

M. RlDDlFORD

principal proleg retractor muscle (PPRM) degenerates during the prepupal period (Fig. 7). This degeneration depends on the prepupal ecdysteroid peak (Weeks and Truman, 1986a; Weeks, 1987), but the commitment to die is dependent on the prewandering surge of ecdysteroid in the absence of JH (Weeks and Truman, 1986b) as is also true of the proleg epidermis itself (Truman et al., 1974). Thus, exogenous JH given systemically to isolated abdomens which are subsequently infused with 20E to mimic this peak

MOT0 R NEURONS

MUSCLES

FIG. 7 Hormonal regulation of metamorphosis of the nervous system and of muscles in Manduca sexfa as illustrated by a larval sensory neuron that changes to activate the pupal gin trap reflex, a motor neuron (PPR) that innervates the principal planta retractor muscle (PPRM) and controls retraction of the larval proleg, and the ventral external oblique muscle (VEO). The change in muscle fibre diameter of both PPRM and VEO is shown. See text for details.


241

resulted in the retention of both the proleg and the muscle, but not the crochets, during the subsequent moult (Weeks and Truman, 1986b). Importantly, localized application of J H over the tip of the proleg during the ecdysteroid release in vivo resulted in the retention of the proleg but did not prevent muscle degeneration (Jacobs and Weeks, 1990), indicating that each tissue responds to ecdysteroids individually. A similar phenomenon was seen with the ventral external oblique muscle which normally dies during adult development; J H only prevented degeneration when given before the prewandering peak of ecdysteroid (Weeks and Truman, 1986b). Thus, just as with the epidermis, the fate of larval muscles at metamorphosis is determined by the small peak of ecdysteroid acting in the absence of JH. In the case of the abdominal intersegmental muscles which die just after eclosion, J H given to the pupa at the outset of adult development can prevent this death (Lockshin and Williams, 1964). Again the critical time for J H action is during the onset of the rise of ecdysteroid for adult development just as in the case of the prevention of the appearance of adult cuticular proteins in the epidermis (Williams, 1961). Localized J H application during this rise also prevents the metamorphosis of larval muscles which grow and reorient to form new attachments on the adult cuticle (J. W. Truman, personal communication). Both growth and reorientation were prevented as well as the formation of discrete motor end plates by the adult motor neurons. Importantly, this effect was seen up to the time that reorientation normally occurs, long after the epidermis became insensitive to J H for adult cuticle formation. Thus, the timing of JH sensitivity of the muscle is later than that of the epidermis. Since muscle reorientation requires information from the epidermis (Willams and Caveney, 1980), it is unclear whether this effect of J H is on the muscle only or also is on the production of these intercellular signals by the epidermis. Development of flight muscles in hemimetabolous insects begins during the final larval instar. In Locustu, both increased muscle growth and the accompanying tracheolization begin late in the instar and are correlated with the pre-ecdysial rise in ecdysteroid (Van den Hondel-Franken, 1982). The commitment of the tracheoblasts to this later differentiation was prevented by J H from adult female corpora allata implanted early in the instar (Van den Hondel-Franken, 1982) or by topical application of methoprene just after ecdysis (Cotton and Anstee, 1990). Also, these treatments reduced both the amount of muscle growth and the production of mitochondria. The same effects on tracheoblasts and mitochondria1 proliferation were seen in the cricket, Teleogryllus oceunicus (Novicki, 1989). In this case after the subsequent moult, the ultrastructural organization was that of the nymph rather than that typically seen in the adult.

L. M. RlDDlFORD

242

3.5

NERVOUS SYSTEM

As with the muscle, the larval nervous system must change at metamorphosis to accommodate the new demands placed by the developing adult. The larval neurons either die or are respecified, and new adult-specific neurons can arise from immature, arrested imaginal cells that have been generated during larval life (Weeks and Levine, 1990). Studies of the hormonal control of these changes have been primarily concerned with the effects of ecdysteroids, and only a few to be discussed below were concerned with the role of J H as well. The larval prolegs of Manduca show a simple withdrawal reflex mediated by mechanosensory planta hairs which make a monosynaptic connection with the motor neurons innervating the proleg retractor muscles (principal (PPR) and accessory (APR) planta retractor motor neurons) (Weeks and Jacobs, 1987; Levine and Weeks, 1990). These hairs and their afferents are normally lost with the proleg at metamorphosis, and the associated motoneurons PPR and APR show a dramatic regression of their dendritic fields. When localized J H was applied to the tips of the prolegs before the commitment rise in ecdysteroid, the hairs and their central projections were retained after pupation, but the PPR motor neurons regressed normally (see below) (Jacobs and Weeks, 1990) so the reflex response of PPRM was lost. Thus, the dendritic regression shown by PPR is not the result of loss of afferents that normally project to the cell. Normally the proleg retractor motoneurons regress as the muscle dies during prepupal development (Weeks and Truman, 1986a) (Fig. 7). Then just after pupation, all PPRs and a subset of APRs die. During adult development, the other APRs which are confined to two segments sprout new dendritic arbors to innervate new abdominal extensor muscles. Just as is true for the planta retractor muscle, these motor neurons are committed to later regression and, in the case of PPR and the posterior APR, death by the prewandering peak of ecdysteroid in the absence of J H (Weeks and Truman, 1986b). These fates are not enacted until they are exposed to the prepupal peak of ecdysteroid. Importantly, regression and death as well as the death of the proleg retractor muscle that they innervate occur at different times and have each been shown to be dependent on a different critical duration of exposure to the hormone, indicating that each of these cellular events is triggered independently (Weeks, 1987; Weeks et al., 1992). Manduca pupae have three sets of abdominal gin traps in which stimulation of mechanosensory hairs causes a reflexive contraction of the abdomen leading to closing of the trap (Levine and Weeks, 1990). This reflex is mediated by surviving larval sensory neurons and intersegmental muscle motor neurons but also involves interneurons, some of which may persist from the larva (Levine and Truman, 1985; Levine et al., 1985, 1989). Although the sensory neurons are the same, their arborization in the central


243

nervous system expands during the larval-pupal transition to mediate a new, pupal-specific reflex (Levine et ul., 1985) (Fig. 7). Localized application of JH at the future site of the pupal gin trap prior to the prewandering peak of ecdysteroid caused the retention of a small patch of larval cuticle and its associated larval hairs with their larval pattern of arborization in an otherwise normal pupa (Levine et al., 1986, 1989). Stimulation of these hairs caused a larval response, whereas stimulation of the pupal hairs on the opposite side of the animal evoked a normal pupal response. Thus, the change in the arborization pattern of these sensory cells is necessary for the pupal response. To determine if this change is sufficient, abdomens were isolated at wandering and 20E was applied topically to cause the formation of a pupal gin trap in an otherwise larval abdomen. Under these conditions, stimulation of the sensory neuron caused a larval motor response. Thus, the sensory neuron must be pupal in order to evoke a pupal response, but other metamorphic changes in the central nervous system are also needed for the reflex to be expressed. The outgrowth of new dendritic arbors of larval motor neurons during respecification in adult development is also dependent on ecdysteroid and can be prevented by J H (Truman and Reiss, 1988). Adult-specific outgrowth of abdominal motoneurons MN-1 and MN-3 begins during the rise in ecdysteroids that initiates adult development. The outgrowth of MN-1 was prevented by JHA (methoprene) only if given early during the rise of ecdysteroid. However, another abdominal motoneuron MN-3 remained sensitive to JH for 2-3 days longer. Thus, different neurons apparently respond to differing levels of ecdysteroid. Whether this differing sensitivity is due to differing levels of ecdysteroid receptors at the outset of the ecdysteroid rise is not known.

3.6

OTHER MORPHOGENETIC ACTIONS

To find simple systems in which to study the morphogenetic actions of JH at the molecular level, there have been in vitro studies on the puffing response in dipteran salivary glands and on various insect cell lines. J H had no effect on the ecdysteroid-induced puffing response of Drosophifu larval salivary glands in vitro, but inhibited the late prepupal response of glands to 20E when given during the prerequisite ecdysteroid-free period for that response (Richards, 1978). The basis for this inhibition has not been elucidated. In salivary glands of the midge, Chironomus tentuns, J H analogues have been reported to activate certain puffs and to prevent some of the normal metamorphic changes (reviewed by Laufer and Borst, 1983), but little further work has been done. In Kc cells (derived from Drosophifa embryos) exposed to ecdysteroid, the JHA methoprene partially inhibited the induced morphological changes

L. M. RlDDlFORD

244

and commitment to proliferative arrest and completely inhibited the appearance of acetylcholinesterase (Cherbas et a f . , 1989). By contrast, methoprene had no effect on the direct induction of the mRNA for the ecdysone-induced protein EIP 28/29. Similar effects were seen with J H I. The inhibition of acetylcholinesterase induction occurred at low levels of methoprene (lop9M) and was not dependent on the ratio of J H to 20E, suggesting that J H was not acting as a competitive inhibitor of the binding of 20E to the regulatory regions of the Eip 28/29 gene. Recently Berger et al. (1992) have shown, in Drosophila S3 cells, that the presence of J H inhibits the ecdysteroid activation of the small heat shock protein (hsp) 27 promoter. When two copies of the 23 bp sequence of the 5' promoter containing the ecdysone response element were fused in tandem to a reporter gene construct, 20E activated expression. Methoprene inhibited this activation in a dose-dependent manner with an EDso between M , but only when the cells were pretreated for 2 h and when and it was present during the ecdysteroid treatment as well. The effect was also seen with J H 111, but not with biologically inactive compounds such as farnesol. Also, methoprene had no effect on the normal heat shock response of the hsp 23 gene. Berger et a f . (1992) therefore conclude that a JH-JH receptor (JH-JHR) complex either prevents or modifies binding of the EcR to the DNA or interferes with subsequent steps in activation of transcription. These findings parallel the effects of J H on ecdysteroidinduced pupal commitment of Manduca larval epidermis in vitro in which pretreatment as well as continuing exposure to JH is also necessary to prevent the action of 20E (Riddiford, 1978). Therefore, the S3 cells seem to be a good model system in which to study the morphogenetic action of J H at the molecular level.

4

Mechanism of JH action

Upon its release from the corpora allata, J H binds to a specific protein in the haemolymph called the J H binding protein (JHBP) and also may bind with low affinity to some of the major haemolymph proteins such as arylphorin (Goodman and Chang, 1985; Goodman, 1990; Kanost et al., 1990). These proteins serve to solubilize and transport this lipophilic molecule and to protect it from degradative enzymes before it reaches its target tissue. Whether these JHBPs are also necessary for delivery into the target tissue is not clear. Preliminary experiments using purified Manduca JHBP with larval epidermis in vitro showed no significant difference in the effective concentration of J H needed to prevent the 20E-induced change to pupal commitment, indicating that this protein was not essential for action on the target tissue.

TABLE 1 Cellular JH binding proteins Tissue

Species

Fraction

Kd (nM)

Epidermis

Drosophila hydei Galleria mellonella Manduca sexta

Cytosolic Cytosolic Nuclear

3.6' 24

Manduca sexta

Nuclear Cytosolic

Locusta migratoria Melanoplus bivattam Leucophaea maderae

Nuclear Nuclear Nuclear Nuclear Cytosolic

0.6 1.85 8-10 2.8 1.7'

JH JH JH JH JH

Heliothis zea Drosophila melanogaster

Cytosolic Cytosolic

4.6 4.5

JH I JH I11

Leucophaea maderae Sarcophaga bullata Sarcophaga bullata Rhodnius prolixus

Cytosolic Cytosolic Membrane Membrane

Silk gland

Galleria mellonella

Cytosolic

15

K, cells

Drosophila melanogaster

Cytosolic Cytosolic

15.6 ND

Fat body

Ovary

I' 4' ND ND

19.1 139 92 6.5

Ligandd

No. of sites Mr

JH I(lOR, 11s) 7.7' J H I(lOR, 11s) ND 10 650 JH I(lOR, 11s) 8 600 IVMA' EBDA, EDHA, MDK ND EDBA, EHDA, MDK ND III(lOR, 11s) 111 111 III(1OR) III(1OR)

200 000 19000 ND ND ND 1.2b ND

X

ND 65 ND ND 29" 38" 80" ND 4-5s ND 65

ND 85"

Reference Klages et al., 1980 Wisniewski et al., 1988 Osir and Riddiford. 1988' Palli et a[., 1990 Braun et al., 1992 Roberts and Jeffries. 1986 Koeppe and Kovalick, 1986 Engelmann et al., 1987 Engelmann et al., 1987; Engelmann, 1990 Muehleisen et al., 1990 Shemshedini et al., 1990

ND

200 241' 440-660 9.5 nMND 1.9' 43 150

JH 111

ND

5CL120

Wisniewski and Kochman, 1984

JH I EFDA

2 500 ND

80 24.6"

Chang et al., 1980 Wane et al., 1989

JH JH JH JH

111 111

I11 I(lOR, 11s)

Koeppe et al., 1981 Van Mellaert et al., 1985 Van Mellaert et a [ . , 1989 Ilenchuk and Davey, 1985

ND, Not determined. Subunit molecular weight as determined by SDS gels. p m o h g protein. Lower affinity binding sites also detected. Racemic unless otherwise specified. Iodovinylmethoprenol. In these studies, 5-10 pg DNA was used per assay rather than the 5-10 ng DNA incorrectly reported in Osir and Riddiford (1988)

'

'

246

L. M. RlDDlFORD

Once at the target, J H has several different modes of action in the cell, depending on the system studied. In the follicle cells of the bloodsucking bug, Rhodnius prolixus (Ilenchuk and Davey, 1985, 1987; Sevala and Davey, 1989), and in the male accessory gland of Drosophila (Yamamoto et al., 1988), JH apparently acts at the membrane level (Table l ) , apparently triggering the phosphatidyl inositol pathway. In fat body, epidermis, ovary, accessory glands, and Drosophila cell lines, intraceliular J H binders are found (for reviews, see Goodman and Chang, 1985; Roberts and Jeffries, 1986; Palli et al., 1991a; Table 1). Although most of these studies consider only one type of action in a particular system, J H may well have pleiotropic effects in a given cell mediated by different receptors. In the stimulation of vitellogenin mRNA production in adult Leucophaea (Engelmann, 1987) and Locusta (Wyatt, 1988, 1990) fat body, J H is thought to act as a steroid hormone and to activate vitellogenin production by induction of vitellogenin mRNA transcription. The long lag between the addition of JH and the appearance of the mRNA, about 12 h in the case of secondary induction, suggests that other factors are required as well. In Locusta protein synthesis is required for this action of JH, but JH is thought to act as a co-regulator on the vitellogenin gene (Wyatt, 1990; Wyatt et al., 1992a) in direct modulation of gene expression. This topic will be covered in detail in Part I1 (Wyatt and Davey, unpublished); here I will concentrate on J H actions in the larva. 4.1

JUVENILE HORMONE BINDING PROTEINS IN THE HAEMOLYMPH

The JHBPs come in two forms (Goodman, 1990; Kanost et al., 1990). In insects that use J H 111, the JHBP is of high molecular weight (near 500 kDa) and shows strong selectivity for binding the natural 10R enantiomer of J H I11 (Kd = 3-9 nM) (deKort and Koopmanschap, 1987). The binding site also apparently requires the epoxide ring and the methyl ester. In Drosophila larvae, the JHBP is also large (400 kDa) and binds racemic J H I11 with nM affinity; both JH I and I1 are weak competitors for the binding (Shemshedini and Wilson, 1988). In Lepidoptera the haemolymph JHBP is small (20-40 kDa) and binds with a Kd of about 100 nM to the natural isomers of J H I and 11; weaker binding is seen with JH I11 (reviewed in Goodman and Chang, 1985; Goodman, 1990). Purification of the Manduca JHBP and subsequent isolation of its cDNA have shown that it is produced by the fat body and has no sequence similarities with other known proteins (Lerro and Prestwich, 1990). Interestingly, the recombinant protein produced in baculovirus from this cDNA binds JH I and JH I1 with Kds of 11 and 42 nM respectively (Touhara et al., 1992). Using the photoaffinity analogue of J H 11, epoxyhomofarnesyl diazoacetate (EHDA), Touhara and Prestwich (1992)


247

have mapped the JH binding site of this protein. They propose that there are disulphide bridges between the N- and C-terminal regions of the molecule which form a hydrophobic pocket into which the lipophilic JH fits.

4.2

INTRACELLULAR JUVENILE HORMONE RECEPTORS

For a hormone to act on a cell, the cell must have receptors for that hormone. These receptors have certain characteristics: (1) they must bind the hormone with high affinity since the hormone concentration is normally low; (2) they must bind the hormone specifically; (3) hormone binding should be saturable and reversible; (4) the tissue and temporal distribution of the receptors should be compatible with the observed actions of the hormone; and ( 5 ) binding of the hormone to the receptor should produce the observed biological effects of the hormone. The traditional approach to the biochemical study of hormone receptors has been to homogenize tissues or cells, isolate various subcellular fractions, and assay these for high affinity, specific hormone binders using radiolabelled hormones. This approach has been complicated in the study of J H receptors due to the hydrophobicity of the hormone, the low specific activity of the available labelled hormones, and the apparent enantiomeric selectivity of the receptor (Klages et al., 1980; Roberts and Jefferies, 1986; Engelmann et al., 1987; Engelmann, 1990; Wyatt et al., 1992a; Braun et al., 1992). However, these types of studies have led to the identification and partial purification of several cellular binding proteins for J H (Table 1). A new approach to isolation and purification of J H receptors (JHR) has been the use of radiolabelled, biologically active photoaffinity analogues of the JHs (Wang et al., 1989; Palli et al., 1990; Prestwich, 1991). Since the analogue is covalently photocrosslinked to the cellular protein(s) to which it is binding, this protein(s) can be tracked by its radiolabel through the purification procedure. 4.2.1

Cytosolic J H binding proteins

Recent isolations of cytosolic binding proteins for J H have yielded a diversity of sizes (Table 1). Using EFDA (10,Il-epoxyfarnesyl diazoacetate), the photoaffinity analogue of J H 111, Wang et al. (1989) isolated a dimeric binding protein with a subunit weight of 24.6 kDa from the cytosol of Drosophila Kc cells (Table 1). In Drosophila larval fat body, Shemshedini et al. (1990) found a 85 kDa cytosolic protein that specifically bound EFDA; these extracts also bound JH I11 with high affinity (see Table 1). A protein with similar properties is also present in larval integument and male accessory glands, but not in adult ovaries or thoracic muscle. The methoprene-tolerant (Met) mutant of Drosophila shows reduced

248

L. M. RlDDlFORD

sensitivity to both J H I11 and methoprene in terms of inhibition of adult differentiation by exogenous hormone and delayed onset of reproduction (Wilson and Fabian, 1986, 1987). This resistance has been linked to reduced binding affinity of a cytosolic J H binder that has been found in both fat body and male accessory glands (Shemshedini et a l . , 1990; Shemshedini and Wilson, 1990). The mutant shows only minor differences in penetration, excretion, tissue sequestration, and metabolism of J H 111, but the cytosolic binding affinity for J H 111 is 6-lO-fold lower than that in the wild type fly. Moreover, Met heterozygotes have two binders, one characteristic of wild type and one of the Met phenotype. By contrast, MetlDf flies have only the J H binder found in Met homozygotes. This genetic and biochemical evidence suggests that the Met locus may encode a cytosolic binder for J H which is critical for its cellular action. Whether this is a JH receptor that upon binding J H enters the nucleus and acts like a steroid hormone receptor complex or whether it is a carrier for this hydrophobic molecule such as the cellular retinoic acid binding protein (Blomhoff et al., 1990) has not yet been determined. A 38 kDa protein that specifically binds the photoaffinity analogues of JH I and J H 11, EBDA (lOR, 11s-epoxybishomofarnesyl diazoacetate) and EHDA (lOR, 11s-epoxyhomofarnesyl diazoacetate), and of methoprene, MDK (7s-methoprene diazoketone), has been found in the cytosol of Manduca 5th instar epidermis (Palli et al., 1990). Binding of each of these analogues can be competed by either the natural JHs or methoprene, indicating that this protein may be a general carrier for the lipophilic JHs and JHAs. It has not been further purified nor its binding affinity determined. A 29 kDa protein in the cytosolic fraction also bound the analogues although its electrophoretic characteristics and binding specificity were similar to the nuclear 29 kDa JH-binding proteins discussed in Section 4.2.2 below, indicating that they may be the same protein. Whether normally it is present in the cytosol or whether this presence is an artefact of the cuclear isolation procedure is not yet known. 4.2.2 Nuclear receptors Although Table 1 shows that high affinity nuclear binders for J H are found in various tissues, little work has been done on their purification. Recently a protein has been found in adult locust fat body nuclear extracts that specifically binds enantomerically pure J H I11 with high affinity (Kd = 0.6 nM) (Braun et al., 1992) (Table 1). This protein will be discussed further in Part I1 (Wyatt and Davey, unpublished). 4.2.2.1 Characteristics of a putative nuclear receptor from larval epidermis. J H readily enters the Manduca larval epidermal cell, and 32% is retained in the nucleus of day 1 5th instar epidermis by two binders with Kds


249

of 6.6 nM and 88 nM for J H I (Osir and Riddiford, 1988) (Table 1). Two nuclear binders of similar affinities for iodovinylmethoprenol were also found. About 10 000 high affinity sites per diploid nucleus appeared to bind these JHs. Interestingly, competition studies indicated that the dodecadienoate JH analogues were unable to compete for binding of the natural JHs and vice versa, indicating that they might be binding to separate molecules or to different sites on the same molecule. A 29 kDa (monomeric weight) nuclear protein of PI 5.8 which specifically bound either EBDA or EHDA was found in both epidermis and fat body of day 1 5th instar larvae (Palli et al., 1990). The same protein non-specifically bound MDK; a second 29 kDa nuclear protein of PI 6.0 bound MDK specifically. The binding of EBDA and EHDA could be competed by the natural hormones, but not by methoprene; that of MDK by methoprene and its derivatives but not by the natural JHs. Thus, these 29 kDa proteins showed similar binding characteristics as found in the earlier studies of Osir and Riddiford (1988) with the isolated epidermal nuclei. The radiolabelled photoaffinity analogues of J H I1 and methoprene were used to purify a 29 kDa nuclear protein from Manduca epidermis that specifically bound the J H I1 analogue EHDA (Prestwich, 1991; Prestwich et al., 1992; Palli et al., 1993). Oligonucleotides prepared from sequences of three peptide fragments were used to obtain a 190 bp cDNA fragment by the polymerase chain reaction (PCR) (Palli et al., 1993). The subsequent isolation and sequencing of the full length cDNA shows that the encoded 29 kDa protein has four small domains of 8-12 amino acids in the Cterminal third of the protein that show 60-75% identity with regions in the bovine interphotoreceptor retinoid binding protein (IRBP) and in human rhodopsin. In IRBP, this identity is in a four-fold protein repeat region of IRBP that is thought to bind fatty acids (Borst et al., 1989); thus, it may be a part of the JH-binding site. A region of 21 amino acids N-terminal to these putative J H binding sites shows 43% identity with the human p68 nuclear protein. The latter protein is an RNA helicase that is found throughout the nucleus in interphase, then localizes to the nucleolus just after cell division (Iggo and Lane, 1989); its function is unknown. Cell extracts containing baculovirus-produced 29 kDa protein specifically bind both J H I and I1 with high affinity (7-11 nM) (Palli et al., 1993), similar to the binding affinity of isolated epidermal nuclei for J H I (Osir and Riddiford, 1988) and only slightly higher than that of the recombinant haemolymph J H binding protein (Touhara et al., 1992). This baculovirusproduced 29 kDa protein also binds EHDA (Palli et al., 1993). However, further purification of the protein is necessary for accurate determination of these binding constants and for a study of the binding site(s) for the natural hormones and the various J H analogues. Immunocytochemical studies with a polyclonal antibody to the 29 kDa protein show that the protein is localized to the nucleus in both epidermis

250

L. M. RlDDlFORD

and fat body of feeding 4th and 5th instar larvae (Palli et al., 1993). Moreover, within the nucleus, staining is confined to subnuclear structures that appear similar to the bismuth-stained nucleolar necklaces found by Tuck and Locke (1985) in Manduca larval epidermis. Studies at the electron microscopic level are presently underway to determine whether the recognized antigen is present in the nucleoli. Initial experiments hybridizing DNA fragments to protein blots had indicated that a 29 kDa nuclear protein specifically bound to both the LCP14 and the LCP16/17 genes with the same developmental pattern as was seen for the 29 kDa JH binding protein (Palli et al., 1990). Yet the deduced sequence of the 29 kDa J H binding protein has no known DNA binding motifs (Palli et al., 1993), and its immunocytochemical nuclear localization is unlike that of the ecdysteroid receptor and MHR3, two known DNA binding proteins (Riddiford, unpublished). Therefore, whether and how this protein interacts with these and other larval structural genes is of great interest. 4.2.2.2 Hormonal regulation of the nuclear 29 k D a J H binding protein The 29 kDa J H binding proteins that bind MDK are present in both 4th and 5th instar epidermis, decrease to low levels during the larval moult, and disappear at wandering in response to 20E in the absence of J H (Palli et al., 1991b) (Fig. 8 ) . They then reappear in the pupal abdominal epidermis. The continued presence of JH proved necessary during both the larval moult and during the intermoult for the maintenance of the normal level of these 29 kDa proteins. Moreover, the reappearance after pupal ecdysis occurred only when J H was present during the prepupal rise in ecdysteroid. Hybridization of the cDNA encoding the 29 kDa protein shows a similar developmental pattern of mRNA expression. The RNA falls to undetectable levels during the latter part of the larval moult although the protein as detected by immunoblotting persists at low levels (Palli et al., 1993) (Fig. 8 ) . By contrast, at the time of wandering and pupal commitment of the epidermis, both RNA and protein disappear. Then just before pupal ecdysis, a trace amount of this mRNA transiently appears. Thus, this 29 kDa nuclear protein is present during the larval growth periods when J H is high as well as at the time that ecdysteroid initiates the larval moult when J H is necessary to prevent metamorphosis. Importantly, it disappears in response to ecdysteroid in the absence of J H at the time of pupal commitment, when the cells become unresponsive to JH. Then it reappears in the pupa when the cells again become responsive to JH. This developmental pattern and the high affinity, specific binding for J H suggests that this 29 kDa protein is critical for the morphogenetic action of JH and thereby should be considered the JH receptor necessary for the hormone’s action.

CELLULAR AND MOLECULAR ACTIONS OF JUVENILE HORMONE HCS

ECDYSIS

0

1

2

3

4

ECDYSIS

0

1

251

WANDERING

2

3

4

5

ECDYSIS ~. ~

6

7

9

A

mRNA

protein

8

7

4

FIG. 8 Developmental pattern of expression of the 29 kDa JH binding nuclear protein from the epidermis of Munducu sextu. mRNA based on Northern and dot blot hybridization using the cDNA encoding the 29 kDa protein (Palli et ul.. 1993); protein based on MDK binding (Palli et ul., 1991b).

4.3

MORPHOGENETIC ACTION OF JH: MODULATION OF ECDYSTEROID ACTION

In its morphogenetic action, J H appears to have no specific action of its own, but rather it alters the molecular responses to ecdysteroid. Thus, the insect always moults, but the presence of J H ensures that the moult will result in the same form and hence that the same genes will be expressed after its completion as before its initiation. To be effective in this maintenance of the status quo, JH must be present in the cell at the time that 20E first enters (Riddiford, 1978; Berger er al., 1992). Since ecdysteroids act at the genomic level, this role of JH as a modulator of ecdysteroid effects is also thought to be manifest through some nuclear action. In Manduca epidermis, one-third of the J H that enters the cell is found in the nucleus, one-third in the 100 000 g pellet (microsomal fraction) with much of the remainder in the cytosolic fraction (Osir and Riddiford, 1988). Therefore, some of its premetamorphic effects could be via membrane actions and second messengers. No studies have yet been done in this realm. How J H modulates ecdysteroid action at the molecular level is the major question still to be answered. Its effect could be direct or indirect. Possible direct effects would be competition of the JH-JH receptor (JH-JHR) complex with the ecdysteroid-EcR (Ec-EcR) complex for binding to the ecdysone response element or the binding of J H at another site that causes interference with subsequent events required for transcriptional activation or

252

L. M. RlDDlFORD

inactivation. Berger et al. (1992) have shown that JH inhibits the 20-HE activation of a gene construct containing only two ecdysone response elements and short flanking sequences (see Section 3.7 above), but whether J H binds to any of these sequences has not been determined. This type of inhibition at or near the ecdysone response elements on the “early” genes encoding transcription factors seems unlikely since J H does not prevent ecdysteroids from initiating and co-ordinating a moult or, as discussed in Section 3 above, from inducing synthesis of proteins which suppress the ongoing transcription of genes that are expressed during the larval intermoult periods. Several of the ecdysteroid-induced transcription factors such as E74 (Thummel et al., 1990), E75 (Segraves and Hogness, 1990; Jindra et al., 1992; Segraves and Woldin, 1992), and MHR3 (Palli et al., 1992) which are involved in the moulting cascade of gene activation and inactivation appear in both larval and pupal moults so may not be influenced by JH. Yet in the case of MHR3, the presence of high J H I in vitro delays the onset of its expression in response to 20E (Palli et al., 1991a); the mechanism of this action is unknown. By contrast, others such as the BR-C proteins first appear at the time of metamorphosis (DiBello et al., 1991; Guay and Guild, 1991; Andres and Thummel, 1992), so their appearance could be either directly or indirectly regulated by JH. Another possible direct effect of J H could be on the target genes of this ecdysteroid-induced cascade. In this case, binding of the JH-JHR complex in the vicinity of a larval-specific gene might prevent its permanent inactivation by interfering with ecdysteroid-induced changes in chromatin structure necessary for that inactivation. In the same way, binding of the JH-JHR complex might stabilize chromatin structure around genes that are not to be expressed until metamorphosis, thus preventing ecdysteroids or ecdysteroid-induced transcription factors access to these genes. However, such an influence on chromatin structure does not require that the JH-JHR complex interact directly with the regulated gene. It could simply influence synthesis and/or activity of one or more of the various nuclear proteins involved in the chromatin changes. The putative JH receptor in Manduca larval epidermis shows weak similarity to an RNA helicase and localizes to substructures in the nucleus that may be associated with the nucleolus (Palli et al., 1993). These data suggest that JH may influence RNA metabolism within the nucleus. Possibilities include effects on post-transcriptional processing of the mRNA, RNA translation, ribosomal RNA production or ribosome assembly. J H may also influence ecdysteroid action indirectly by regulating the ecdysteroid receptor level in cells and thus their responses to the changing ecdysteroid titre. The larval motor neurons in Manduca show no ecdysteroid receptors during larval life except trace amounts just before each ecdysis (Riddiford and Truman, 1993). Then all, irrespective of whether they are to be remodelled or to die during metamorphosis, show high levels at the time


253

of pupal commitment. Similar increases in levels of EcR could be induced in isolated abdomens by infusion of 20E, but was prevented by the simultaneous presence of the J H A methoprene (M. Renucci and J. W. Truman, personal communication). In the epidermis of Manduca, ecdysteroid receptors (based on both mRNA and immunocytochemical studies) are present at low levels in larval epidermis at all times; yet there is also a significant and transient increase at the time of pupal commitment (Riddiford and Truman, 1993; Riddiford, Maves, Newitt and Palli, unpublished). Whether this increase can be prevented by JH is currently being studied. Whether there is also a qualitative change in ecdysteroid receptor isoform at metamorphosis in Munduca is unknown. In Drosophifa the larval motor neurons show no ecdysteroid receptors until midway in the final larval instar at which time they all acquire high levels of the B1 isoform of the receptor by pupariation (Truman et al., 1993). Later during pupal development they lose this form, and a subset that is fated to die at the end of metamorphosis acquires high levels of the A isoform. Whether this change in receptor isoform is dependent on the absence of J H has not been studied. An analogous situation exists in amphibian metamorphosis where thyroid hormone induces metamorphosis and prolactin inhibits this action (Dent, 1988; Tata et al., 1991). Normally during larval life, the a isoform of the thyroid hormone receptor is present; then at metamorphic climax the p isoform becomes predominant (Yaoita and Brown, 1990; Kawahara ef al., 1991). Prolactin prevents the appearance of this fl isoform (Baker and Tata, 1992). Presumably then the genes that are regulated by the new isoform cannot be activated or repressed so that metamorphosis cannot proceed. Thus, the action of J H in preventing metamorphosis of insects could be similar.

5 Juvenile hormone analogues as insect growth regulators

When first reporting the extraction of J H from a natural source, Williams (1956) suggested the use of this hormone as a pesticide, since it could be absorbed through an insect’s cuticle and prevent normal development, and would presumably be refractory to development of resistance. Later, the discovery of the “paper factor” (Slama and Williams, 1965), a J H analogue active only in the hemipteran family Pyrrhocoridae, indicated the potential of such analogues in the selective control of insects. Thus, they were regarded as “third generation pesticides” (Williams, 1967), and programmes directed toward the synthesis of J H analogues that would combine high biological activity with chemical and biological stability were set in motion. From among the hundreds of compounds synthesized, many were active as

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“insect growth regulators” (IGRs) and a few (Fig. l b ) have found, or currently show promise for, practical use in the control of pest species (for reviews, see Jacobson et a f . , 1972; Slama et a[., 1974; Staal, 1975; Henrick et af., 1976; Sehnal, 1983; Retnakaran et al., 1985; Hatakoshi and Nakayama, 1987; Langley et al., 1990). Some of these IGRs are also widely used by researchers as J H agonists, in order to avoid the rapid inactivation of experimentally applied natural J H . Whereas JHAs such as methoprene and hydroprene are structurally related to J H (Fig. 1) and were designed for resistance to biological degradation as well as chemical stability (Henrick et a f . , 1973), a surprising finding was the activity of many substances that differ widely in chemical structure from JH, and frequently possess aromatic functions. The JHAs with activities among the highest yet reported, fenoxycarb (Roche/Maag; Dorn et af., 1981; Masner et a f . , 1981) and pyriproxyfen (Sumitomo Chemical Co.; Hatakoshi et a f . , 1986, 1987) (Fig. lb), are aromatic compounds with little apparent resemblance to JH. While their stability is clearly one aspect of the high activity of these substances, recognition by a specific receptor site and ability to activate the responding cellular system are also required. The biological effects of various JHAs are remarkably similar to those of the natural JHs, allowance being made for differences in effective dose and stability. This suggests that they share common modes of action and thus one would expect a common receptor. Yet the diversity of structures of the active analogues and the strict steric requirements for biological activity of the natural JHs (Kindle et af., 1989; Sakurai et a f . , 1990) as well as the apparent inability of low amounts of active analogues to compete for binding of the natural JHs in crude cytosolic or nuclear extracts (Osir and Riddiford, 1988; Goodman, 1990; Braun et al., 1992) are paradoxical. This problem can only be resolved by studies on binding and competition with purified receptors.

6 Conclusions

Most revolutionary for an understanding of how hormones control moulting and metamorphosis are the recent revelations from studies on the ecdysteroid receptor, both its isoforms and its heterodimerization requirements for activity, and the ecdysteroid-induced transcription factors in Drosophila. Whether or not structural genes in a particular tissue are regulated by ecdysteroid is first of all dependent on whether or not that tissue has ecdysteroid receptors at that time. If receptors are present, then the ecdysteroid-induced transcription factors will be induced, the form depending on the concentration of ecdysteroid and possibly on the tissue.


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Then these factors, and/or possibly ones that they in turn gencratc as part of the ecdysteroid-induced cascade, act in concert with tissue-specific factors and possibly with the ecdysteroid-ccdysteroid receptor complex itself to regulate the structural genes. In Drosophilu the ecdysteroid receptor has at least three isoforms which differ only in the N-terminal region that is thought to be important in the transactivation of the gene. Therefore, different isoforms may regulate different genes and/or the same genes differently. Within a tissue these receptor isoforms appear to change as development proceeds. Thus. a polymorphic cell such as the epidermis may show a changing array of isoforms as it progresses through metamorphosis. Most of the known ecdysteroid-induced transcription factors appear to be activated in every moult, irrespective of whether it is larval, pupal, or adult. At least one appears only at metamorphosis. These then must regulate different stage-dependent genes in a polymorphic tissue and initiate differentiation in the various imaginal precursor cells and discs. In addition to possible new isoforms of the ecdysteroid receptor, the transcription factors that they induce are found in multiple forms generated by alternative splicing or alternative promoters. This variety provides a multitude of combinatorial possibilities to regulate a particular gene. How J H fits into this picture is still a mystery. Does the JH-JH receptor complex interact directly with the active larval structural genes themselves and/or with the inactive pupal and adult genes or through protein-protein interaction with other transcription factors? O r does it simply influence the type of ecdysteroid receptor and/or specific transcription factor present, and, if so, how? Does J H affect RNA metabolism, influencing transcription, translation or the presence of possible regulatory RNAs? The experiments to provide answers to these types of questions are now feasible and should provide a firm understanding of the molecular basis of the morphogenetic action of juvenile hormone.

Acknowledgements I thank Dr Gerard Wyatt for fruitful discussions on the scope of this review and for constructive criticisms on the manuscript; Dr James Truman for many helpful discussions, a critical reading of the manuscript, and help in preparation of the figures; Dr Kiyoshi Hiruma, Dr Subba Reddy Palli and Dr Rosalie Langelan for their contributions to the unpublished work cited herein and comments on the manuscript; Dr Hiruma for help with the figures; and Dr David Schooley for providing Fig. l a . The cited unpublished work from my laboratory was supported by NSF DCB90-05202 and DCB9106463, NlH A112459 and NS29971, and USDA 91-37302-6207.

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Mechanism of Action of Bacillus thuringiensis lnsecticidal 6- E ndotox ins Barbara H. Knowles Department of Zoology, University of Cambridge, Downing Street, Cambridge, CB2 3EJ, UK

1 Introduction 275 1. I What is Bacillus rhuringiensis? 276 1.2 Classification of Bt toxins 277 1.3 What is Bt used for? 278 2 The structure of Bf toxins 279 2.1 The structure of Cry toxins 279 2.2 The structure of Cyt toxins 280 3 The insect gut 282 3.1 Lepidoptera 282 3.2 Diptera 284 3.3 Coleoptera 284 4 Mechanism of action 285 4.1 Solubility 286 4.2 Activation 287 4.3 The peritrophic membrane 288 4.4 Receptors 288 4.5 Pore formation 291 4.6 Cell lysis 291 5 Models for the mechanism of pore formation 5.1 The “penknife” model 296 5.2 The “umbrella” model 298 6 Conclusions and future prospects 298 Acknowledgements 299 References 299

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1 introduction

Growing public concern about the use of chemical insecticides has led to increasing interest in the biological alternatives. Bacillus thuringiensis (Br). a family of bacteria which make insecticidal proteins, accounts for 9045% of the insect biocontrol market. There have been a number of recent reviews of the use of Bt as a biological insecticide (Peferoen, 1091; Feitelson et G I . , ADVANCES IN INSECT PHYSIOLOGY VOI. 24 ISBN IbI 2-II?J??J-9

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1992; Lambert and Peferoen, 1992), the mode of action of the insecticidal toxins (Chilcott et al., 1990; Ellar, 1990; Wolfersberger, 1990a; English and Slatin, 1992; Gill et al., 1992; Knowles and Dow, 1993) and the diversity and expression of toxin genes (Hofte and Whiteley, 1989). This chapter is therefore not intended as an exhaustive review of the recent literature. I will discuss how the recently solved X-ray crystal structure of one Bt toxin has allowed us to model the mode of action of a whole family of related protein toxins, and how a knowledge of the physiology of the insect targets of Bt toxins is needed to further our understanding of the toxic mechanism. However, I will begin with an overview of the biology of Bt.

1.1

WHAT IS BACILLUS T H U R I N G I E N S I S ?

Bacillus thuringiensis (Bt) is the name given to a family of bacteria found throughout the world. It was first discovered in Japan in 1901, killing silkworms in a silk farm, where it was considered a pest (Ishiwata, 1901). About ten years later another strain was found in Germany, doing a useful job of killing grain moth larvae in stored grain (Berliner, 1915). Since the latter strain was discovered in the province of Thuringen it was named Bacillus thuringiensis, a name now given to a large family of bacteria producing insecticidal parasporal crystals. It was believed for many years that the insecticidal spectrum of Bt was confined to the larvae of Lepidoptera, until screening programmes revealed new strains active against Diptera (Goldberg and Margalit, 1977) and Coleoptera (Krieg et al., 1983). Bt is a rod-shaped gram positive bacterium with a fascinating biology. When nutrients are abundant it grows vegetatively, but when its food supply runs short it makes a dormant spore and one or more large crystalline parasporal inclusions. These crystals contain insecticidal proteins, called bendotoxins, which are lethal when eaten by a susceptible insect. Since the crystals contain more than 30% of the protein made by Bt at a time when it is preparing to resist starvation, one would expect them to play an important role in the survival of the organism. It is thought that the b-endotoxins provide Bt with a selective advantage over other spore-formers (Ellar, 1990): the guts of susceptible insects usually have a high pH, which prevents germination of ingested spores, and a rapid transit time for ingested material. The 6-endotoxins cause paralysis of the gut, allowing the spores to be retained, and break down the gut wall of the insect. The gut contents mix with the blood, thus lowering the pH and providing nutrients to trigger germination of ingested Bt spores. The dead insect then acts as a food source for vegetative bacterial growth, before starvation again leads to sporulation. There are some questions about whether insects are the natural target for Bt toxins (see below) but one piece of circumstantial evidence suggests that Bt has co-evolved with insects: compared with related bacteria,

MECHANISM OF ACTION OF BACILLUS THURINGIENSIS

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Bt is highly resistant to both cellular and humoral components of the insect immune system (Dalhammar and Steiner, 1984; Bornan and Hultmark, 1987). The only other bacteria known to have similar defences against the insect immune response are those that are obligate or occasional insect pathogens. Screening programmes have shown that Bt can be found almost everywhere, in habitats ranging from the Arctic to the tropics (Martin and Travers, 1989). Although most of the original strains were discovered associated with dead insects (Heimpel and Angus, 1960), in insect breeding environments or in stored product residues where insects were abundant (Meadows et al., 1992), Bt does not have an obligate relationship with insects (Dulmage and Aizawa, 1982). Indeed soil screening revealed Bt strains in areas with few or no insects (Martin and Travers, 1989). This raised questions about the ecology of Bt:can it really be considered to be a natural insect pathogen if it is so often found in the soil, while the insects that it is known to kill (mainly larvae of leaf-eating Lepidoptera and Coleoptera, and aquatic larvae of Diptera) are usually present in different environments? The recent finding that Bt is abundant on the surface of leaves from a wide range of deciduous and coniferous trees (Smith and Couche, 1991), and the inference that Bt might be a natural plant dweller, may go some way towards answering this question. However, a proportion of Bt strains produce crystalline protein inclusions which are not toxic to any insect tested (Ohba and Aizawa, 1986; Martin and Travers, 1989; Meadows et al., 1992). Although only a few insects were tested in the latter reports, and the insects screened were those of economic importance or which were easy to rear, it remains possible that there are non-insect organisms, maybe soil dwellers, which are the target of these apparently non-toxic Bt strains (Lambert and Peferoen, 1992). Some Bt strains have been reported to be active against nematodes (Bone et al., 1988; Feitelson et al., 1992). A recently cloned &endotoxin had silent activity against Coleoptera, being non-toxic when fed to beetle larvae unless pre-activated in vitro (Lambert et al., 1992). The natural target of such proteins remains to be identified.

1.2

CLASSIFICATION OF BI TOXINS

Classification has been a problem, with thousands of Bt strains isolated-ne estimate suggests 40 000 in public and private collections (Lambert and Peferoen, 1992). The most recent and helpful scheme, summarized in Table 1, classifies Bt toxins on the basis of DNA homology of the toxin genes and insecticidal activity spectrum of the proteins (Hofte and Whiteley , 1989). Under this scheme most Bt toxins are classified as “Cry”: proteins which are highly specific for insect cells both in vivo and in vitro. A few Bt crystal toxins have a broad spectrum cytolytic activity in v i m , in addition to a

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specific action on dipteran insects in vivo, and are called “Cyt”. The 6endotoxins range in size from 140 to 27 kDa. Each strain of Bt may synthesize more than one class of a-endotoxin. Inevitably there are some Bt 6-endotoxins that do not fit conveniently into this classification scheme. Some of these are indicated in Table 1. TABLE 1 Classification of Bt toxins” Class Cry1 Cry11 Cry111 CryIVA, B CryIVC, D CYt a

Mr Wa)

Specificity

Crystal shape

Exceptions

130-140 70 70 130 70 27

Lepidoptera LepidopterdDiptera Coleoptera Diptera Diptera DipterdCytolytic

Bipyramid Cuboid Rhomboid Bipyramid Bar shaped Amorphous

CryIB’ CryIIB‘ CryIIICd

Based on the classification scheme of Hiifte and Whiteley (1989). CryIB family has a member (originally designated CryV) that kills both Lepidoptera and Coleoptera (Tailor et al., 1992). CryIIB is non-toxic to Diptera (Widner and Whiteley, 1989). CryIIlC is 130 kDa and has cryptic coleopteran activity (Lambert el al., 1992).

1.3

WHAT IS ~t USED FOR?

Since the 1960s Bt has been widely used as a commercial insecticide to kill agricultural and forestry pests (reviewed by Feitelson et al. (1992) and Lambert and Peferoen (1992)) and in a very successful World Health Organization programme in West Africa to control the simuliid blackfly which are vectors of Onchocerciasis (river blindness) (Guillet et al., 1990). The advantages of Bt as a “green” insecticide are its specificity (it is nontoxic to mammals, birds and even to most beneficial insects, such as bees) and the slow rate of development of resistance to Bt by insects. It was for the latter reason that the Onchocerciasis programme was so successful: the blackfly larvae developed resistance to conventional chemical pesticides in as little as 3 months, but to date have not become resistant to Bt after more than 10 years of intensive use (Guillet et al., 1990). Some resistance of lepidopteran pests has been seen in the field and will be discussed in Section 4.4.3. The narrow spectrum of activity of Bt toxins makes them much faster and cheaper to register than new chemical pesticides. Ironically some of the disadvantages of Bt as an insecticide are the same as its advantages: specificity to a narrow range of insects and lack of persistence in the field. These problems are being addressed by a range of


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biotechnological approaches, such as cloning and expression of toxin genes in plants, reviewed by Peferoen (1991).

2 The structure of Bt toxins

2.1

THE STRUCTURE OF CRY TOXINS

As more Bt toxin genes were cloned and sequenced it became apparent that Cry toxins were a family of related proteins, with regions of identity or high similarity interspersed with variable regions (Hofte and Whiteley, 1989; Hodgman and Ellar, 1990). It has been known for many years that most Bt toxins are synthesized as inactive protoxins which are activated by proteolysis in the insect gut (Lecadet and Martouret, 1965). Biochemical and genetic methods revealed that activation of the 130-140 kDa protoxins occurred by removal of the C-terminal half of the protein, leaving an active toxin of around 60 kDa (reviewed by Hofte and Whiteley (1989) and see Section 4.2). Alignment of the sequences of Cry toxins with different specificity revealed five highly conserved sequence blocks in the active Nterminal half of the proteins, separated by hypervariable sequences (Hofte and Whiteley, 1989; Fig. 1). On the basis of sequence comparisons Hodgman and Ellar (1990) proposed that two or more putative amphipathic a-helices in the N-terminal half of the activated 60-65 kDa proteins were responsible for cytolytic activity while the variable C-terminal half of the active toxin was the specificity-determining region, with these two putative functional domains responsible for the proposed two-step mechanism of action of the toxins: receptor binding followed by insertion into the target membrane to form a pore (Knowles and Ellar, 1987). Biochemical and biophysical evidence supported the hypothesis that the active portion of Bt toxins consisted of two folding domains, the N-terminal half of the 60-65 kDa toxin predominantly a-helical and the C-terminal half mainly psheet (Choma et al., 1990; Convents er al., 1990). Perhaps surprisingly, the C-terminal half of the protoxin, which is removed during activation, is even more highly conserved than the active N-terminal half of the protein (Hofte and Whiteley, 1989). This region may be involved in crystallization and in the insolubility of the protein (see Section 4.1). Until recently modelling of toxin structure was largely based on hypothesis, but now the X-ray crystal structure of a Bt beetle toxin (CryIIIA) has been solved by Li er al. (1991). CryIIIA is a 73-kDa protein, whose sequence can be aligned with the active N-terminal half of the larger Cry protoxins. The structure of an active 67-kDa portion of CryIIIA is made up of three distinct domains (Fig. 2): domain I is a bundle of seven amphipathic and hydrophobic a-helices, domain I1 is three antiparallel P-sheets, and domain I11 is a tightly packed P-sandwich in which the

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IIA IVA IIIA IVB

N-

-C

FIG. 1 General features of Cry toxins. This linear representation of Cry toxins shows the three domains of the CryIIIA structure (labelled I , 11, 111, separated by bold vertical lines) (Li et al., 1991) superimposed on an alignment of the active Nterminal half of Cry toxins, extending from amino acids 1-616 in the CryIA(a) sequence. Conserved sequence blocks (Hofte and Whiteley, 1989) are stippled. Arrows mark proteolytic cleavage sites: most Bt toxins are cleaved at around amino acid 29 (left hand arrow); additional proteolytic nicking has been seen in vitro at the indicated positions for CryIIA, IIIA, IVA and IVB (see Section 4.2).

C-terminus is buried. Sequence alignment of other Cry toxins shows that the five conserved regions fall within the internal parts of the CryIIIA structure, in particular at the interfaces of the three domains, suggesting that all Bt Cry toxins have the same basic skeleton (Li et af., 1991). The two folding domains proposed from proteolysis and unfolding studies (Choma et al., 1990; Convents et af., 1990) correspond to (1) the a-helical domain I, and (2) domains I1 and I11 (mainly fi-sheet) together as a single unit. The most exciting feature of this structure is that it immediately suggests what the functional domains of the toxin might be: the amphipathic helices of domain I are good candidates for a membrane insertion/toxicity domain, the variable loops of domain I1 (reminiscent of the antigen binding region of immunoglobulins) are likely to be involved in receptor binding and domain I11 could protect the C-terminus of the active toxin from further proteolysis in the insect gut (Li et af., 1991). The recently solved structure of another bacterial protein toxin, diphtheria toxin, is a second example of a protein with three functional domains (Choe et al., 1992). In this case there is an ADP-ribosylating catalytic domain, a membrane insertion domain (nine ahelices) and a receptor binding domain (two fi-sheets) (Choe et d . , 1992).

2.2

THE STRUCTURE OF CYT TOXINS

Even before the structure of Cry IIIA was solved by X-ray crystallography, comparison of the sequences of the large family of Cry toxins allowed some informed speculation about the toxin structure and the functional domains within the proteins (Convents et af., 1990; Hodgman and Ellar, 1990). The same process was more difficult for the CytA toxin because only two genes had been sequenced, and these showed only a single amino acid difference


281

FIG. 2 A schematic ribbon diagram of the CryIIIA structure (Li ei al., 1991). Domain 1, the putative membrane insertion domain, is a seven helix bundle (left); domain 11, the putative receptor binding region, is an assembly of three P-sheets (lower right); domain 111 is a &sandwich in which the C-terminus is buried (upper right).

(Waalwijk et al., 1985; Earp and Ellar, 1987). Modelling of the secondary structure of CytA was carried out on the basis of computer predictions and site-directed mutagenesis (Ward et al., 1988). A topological scheme for the structure of CytA in a membrane was proposed using secondary structure and hydrophobicity predictions, and proteolytic cleavage patterns (Chilcott et af., 1990). Preliminary X-ray diffraction analysis of CytA suggested that the protein may contain parallel or antiparallel helical bundles (McPherson et al., 1987). The recent publication of the sequence of a second member of the Cyt family, CytB (Koni and Ellar, 1993), enables us to refine the structural model. CytA and CytB show 39% identity of amino acid sequence and 70% similarity, with a pattern of highly conserved regions separated by variable regions as in the Cry family. Structural predictions based on computer algorithms and alignment of CytA and CytB sequences suggested that both

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proteins may have five short fl-strands and six longer a-helices, at least four of which are predicted to be amphipathic and of sufficient length to span a membrane (Koni and Ellar, 1993). Although immunological crossreactivity between Cyt and CryIVD has been reported (Held et al., 1990) and a weak similarity was apparent at the sequence level (Koni and Ellar, 1993) there is not enough information yet to support the proposal that there may be a structural similarity with the Cry family. Nevertheless, in vitro assays suggest a similar mode of action and pore size for Cry and Cyt toxins (Knowles and Ellar, 1987; Drobniewski and Ellar, 1988).

3 The insect gut

The biological target for Bt &endotoxins is the midgut epithelium of susceptible insects. To understand the toxic mechanism we must first find out something about the nature of this target. Most of the research on the mode of action of Bt toxins has centred on toxins active against lepidopteran larvae, but dipteran and coleopteran larvae are also killed by Bt. The structure and function of the insect midgut has been comprehensively reviewed (Dow, 1986) but I will recap the features salient to Bt action in the following sections.

3.1

LEPIDOPTERA

Some lepidopteran larvae can increase in weight by three orders of magnitude in 2 weeks because they spend most of their time feeding. The midgut, being the site of most of the digestion and absorption of food, is therefore one of the most important parts of the larva’s physiology: half of the weight of a lepidopteran larva is its gut and most of this is the midgut (Dow, 1986). Lepidopteran larvae are characterized by a high blood ratio of K+:Na+ (Sutcliffe, 1963) and a very high ambient alkalinity in the gut, exceeding pH 12 in some insects (Dow, 1984, 1986). Many species of lepidopteran larvae are killed by Bt toxins from groups Cry1 and CryII. The midgut of lepidopteran larvae is a simple tube made of one layer of cells resting on a basal membrane. The two major cell types (cartooned in Fig. 3) are columnar cells, with an apical brush border of microvilli, and a unique goblet cell containing a large cavity (filled with a flocculent matrix of sulphated glycosaminoglycans) which is joined to the apical surface by a complex interdigitated valve (Flower and Filshie, 1976; Cioffi, 1979; DOW, 1986; Moffett and Koch, 1992). The epithelial cells are joined primarily by septate junctions and gap junctions (Lane and Skaer, 1980; Lane et al., 1989). The gap junctions provide electrical and chemical coupling between

283


K+ aa

5

4’

5

FIG. 3 Schematic view of the two principal cell types of the lepidopteran larval midgut: the columnar cell (left) and the goblet cell (right). The apical (gut lumen) side is at the top of the diagram. The cells sit o n a basement membrane and are connected by septate junctions and gap junctions. Major ion movements are indicated with arrows. The main pumps and channels are numbered as follows: (1) H+-ATPase; (2) K+/nHf exchanger; ( 3 ) Kf/amino acid co-transporter; (4) gap junction; ( 5 ) basal K + entry through channels and a pump; (6) K + leaving the goblet cavity through the apical valve.

the goblet and columnar cells (Moffett and Koch, 1988b; Dow and Peacock, 1989). A peritrophic membrane, mainly composed of chitin and glycoproteins, lines the gut, separating the gut contents from the epithelium (Richards and Richards, 1977; Adang and Spence, 1981). The physiology and function of the lepidopteran gut is dominated by the vigorous electrogenic transport of Kf from the blood side to the lumen side (Harvey and Nedergaard, 1964). The “K’-pump” is located on the apical membrane of the goblet cell (Dow et al., 1984; Moffett and Koch, 1988b; Moffett and Koch, 1988a; Wieczorek et al., 1989) and is now known to consist of a proton pump (of the VATPase family) in parallel with a Hf/Kf exchanger. This has the net effect of a “futile” cycle of Hf fuelling an electrogenic flux of K+ from the goblet

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cell cytoplasm to the goblet cavity and hence to the gut lumen (Wieczorek et al., 1991) (Fig. 3). The vigorous activity of this pump maintains a potential across the goblet cell apical membrane of up to 270 mV (Dow and Peacock, 1989) leading to a transepithelial potential difference of over 150 mV. The high lumen pH is maintained, at least in part, by a simple Nernstian distribution of protons across the apical membrane (Dow, 1992). Na+/K+ ATPase activity has never been identified despite diligent searching (Anstee and Bowler, 1979; Harvey et al., 1983); and, although there is evidence that the basal entry of K+ cannot be entirely passive (Moffett and Koch, 1988a) and there is active transport of most other ions (Chamberlin, 1990), it is generally agreed that the potentials both across the epithelium as a whole and between the various cellular compartments are determined mainly by the activity of the electrogenic V-ATPase in the goblet cavity membrane. VATPases and their role in the physiology of the lepidopteran gut and other systems are comprehensively reviewed in a volumne edited by Harvey and Nelson (1992).

3.2

DIPTERA

Bt toxins from groups CryIV, Cry11 and Cyt are toxic to some dipteran larvae from the suborder Nematocera, including filter-feeding aquatic larvae of certain mosquitoes and simuliid blackflies (Goldberg and Margalit, 1977; Lacey and Federici, 1979) and the root-feeding tipulid leatherjacket (Waalwijk et al., 1992). Dipteran gut physiology is reviewed by Dow (1986). Like the lepidopteran larvae described above, dipteran larvae sensitive to Bt toxins have a very high pH in their gut (Dadd, 1975; Lacey and Federici, 1979) and utilize mainly serine proteases (Kunz, 1978). Gut pH in mosquito larvae falls rapidly if the larva is chilled, narcotized or manipulated, implying that the high pH is maintained by an active metabolic process carried out by the midgut epithelial cells (Dadd, 1976), although the mechanism of this process is unknown. The gut of mosquito larvae is a simple cuboidal epithelium, with no goblet cells. Unlike Lepidoptera and many plant-feeding Coleoptera, dipteran larvae have a very low K+:Na+ ratio in the blood (Sutcliffe, 1963). Recent reports show that formulations of Bt spores and crystals are toxic to mushroom flies of Lycoriella spp. (Diptera: Sciaridae) (White and Jarrett, 1990; Keil, 1991). It is not yet clear whether the toxic agent is a &endotoxin.

3.3 COLEOPTERA The physiology of coleopteran guts is comprehensively reviewed by Crowson (1981). Like mosquito larvae, coleopteran larvae possess a simple


285

cuboidal epithelium, and lack the specialized goblet cells found in Lepidoptera (Bravo et al., 1992b). Bt toxins kill beetles from two main suborders: crysomelid leaf-eating beetles and scarabaeid white grubs which feed on roots. These two groups have very different gut physiologies. Unlike the dipteran and lepidopteran targets for Bt, crysomelid beetle larvae have a gut pH that is neutral or slightly acidic (Crowson, 1981; Weltens et a f . , 1992) and proteases of the cysteine and aspartate type (Murdock et af., 1987; Thie and Houseman, 1990). These larvae, for example the Colorado potato beetle, Leptinotarsa decemfineata, and the Southern corn rootworm, Diabrotica undecimpunctata howardi, are killed by CryIII toxins (Krieg et al., 1983; Rupar et al., 1991). Several Bt strains, containing toxins which have not yet been characterized, kill scarabaeid larvae such as the Japanese beetle, Popilfia japonica (Sharpe, 1976; Ohba et al., 1992), and the New Zealand grass grub, Costelytra zeafandica (C. N. Chilcott, personal communication). These larvae typically have a high midgut pH (Crowson, 1981) and proteases of the serine type (Christeller et a/., 1989). Their guts therefore present an environment more similar to lepidopteran and dipteran larvae than to crysomelid larvae. Both crysomelid and scarabaeid larvae have the high blood K+:Na+ ratio seen in Lepidoptera, a feature considered to be an adaptation to a plant diet (Sutcliffe, 1963).

4

Mechanism of action

It is widely accepted that the primary action of Bf toxins is to lyse midgut epithelial cells in the target insect, and that the toxins act from the outside of the cell, inserting into the plasma membrane but not entering the cytoplasm (reviewed by Luthy and Ebersold, 1981; Gill et al., 1992). The time course for histological and biochemical changes in lepidopteran insects fed on Bt toxins was first studied in the 1950s and 1960s by Heimpel and Angus (1960) and is summarized in Table 2. The earliest histological effect is a swelling and blebbing of the columnar cell microvilli, followed by various changes to internal membrane structures, cell swelling and eventual lysis of the columnar cells (Endo and Nishiitsutsuji-Uwo, 1980; Luthy and Ebersold, 1981; Percy and Fast, 1983; Lane et a f . , 1989; Bravo et a / . , 1992b). The effect on goblet cells is slower (Bravo et a / . , 1992b): the goblet cell cytoplasm condenses while the goblet cavity swells (Endo and NishiitsutsujiUwo, 1980; Gupta et af., 1985). The onset of the first pathological symptoms is remarkably rapid, especially considering that the protoxins must reach the midgut, dissolve and be proteolytically activated. The basic histopathology of Bt toxins on dipteran and coleopteran larvae is similar to that described for Lepidoptera (de Barjac, 1978; Lacey and Federici, 1979; Bravo et a / . ,

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1992b), although in one coleopteran insect damage to the apical microvilli was not observed (Bauer and Pankratz, 1992). Owing to the small size of the larvae there have been very few studies on the associated biochemical changes in Diptera and Coleoptera. TABLE 2 Time course of Bt pathology in lepidopteran larvae in vivo" 1-5 min

Increased glucose uptake, gut cells First histopathology, columnar cells

5-10 min

Midgut paralysis Cessation of feeding Apical membrane permeable to dyes Swelling of columnar cells Blebbing of columnar cell microvilli First histopathology, goblet cells Increased blood pH, decreased lumen p H

1C-30 min

Increased K' turnover, gut cells Increased blood K + Decreased glucose and leucine transport to blood General metabolic breakdown of gut cells

3 M O min

Cells lyse and slough from basement membrane General paralysis (1-7 h)

1-3 days

Death (starvation, septicaemia)

' This table combines data from numerous studies conducted on Bombyx mori and Manduca sexta, Type I insects (Heimpel and Angus, 1960). Papers cited: Heimpel and Angus, 1959, 1960; Fast and Donaghue, 1971; Fast and Morrison, 1972; Ebersold er al., 1978; Luthy and Ebersold, 1981; Percy and Fast, 1983).

Despite the rapid onset of pathological symptoms, the intoxicated insect usually does not die for several days after ingesting a lethal dose of Bt toxins. Fortunately for its utility as a commercial insecticide, one of the earliest symptoms of Bt action is inhibition of feeding. Death usually results from starvation, or septicaemia as bacteria multiply in the haemocoel. In the following sections I will consider the action of the toxins in order of the proposed steps in their mechanism.

4.1

SOLUBILITY

Most Bt crystal b-endotoxins are insoluble except at high pH. Insolubility is partly conferred by interchain disulphide bonds (Bietlot et al., 1990). The cysteine residues in most Cry toxins are located in the first few amino acids at the N-terminus and in the C-terminal half of the molecule (Hofte and Whiteley, 1989): these are the regions that are removed during activation (see Section 4.2) and the cysteines are not required for toxicity (Choma and


287

Kaplan, 1992). As detailed in Section 3, the midguts of most sensitive insects are characterized by high pH and reducing conditions, suitable for rapid dissolution of 6-endotoxins. High pH may be required for optimal binding as well as solubility (Gringorten et al., 1992). The CryIIIA toxin structure reveals that there are no disulphide bonds in this protein but the crystal packing is stabilized by four interchain salt bridges (Li et al., 1991). These might account for the solubility properties of this toxin: it is only soluble above pH 10 and below pH 4 (Koller ef af., 1992) or in high concentrations of chaotropic salts such as NaBr (Bernhard, 1986). The CryIIIA toxin kills crysomelid beetles, whose gut pH is only slightly acidic (Crowson, 1981; Weltens et al., 1992) so we must presume that in this case other factors in the gut contents promote solubility (Slaney et al., 1992). It is interesting to note that the proteolytically activated CryIA(c) toxin, which has lost all of its cysteines, is now, like CryIIIA, soluble below pH 3.5 as well as above pH 9.5 (Bietlot et al., 1989).

4.2

ACTIVATION

Many Bt &endotoxins are protoxins, activated by insect gut proteases. These typically cleave some 500 amino acids from the C-terminus of CryI protoxins and 28 amino acids from the N-terminus, leaving a 65-55 kDa protease-resistant active core comprising the N-terminal half of the protoxin. The C-terminal half is removed by sequential cleavage of 10-35 kDa fragments which are themselves rapidly degraded into peptides (Chestukhina et al., 1982; Choma et af., 1990). The smallest toxic fragments of Cry toxins identified by genetic deletion experiments are larger than those obtained by proteolysis of full length proteins, possibly because truncated gene products do not fold correctly in the absence of elements which can be removed after folding. Alignment of the amino acid sequences of Cry proteins (Hofte and Whiteley, 1989; Hodgman and Ellar, 1990) revealed that the 70 kDa proteins (CryII, Cry111 and CryIVD) could be considered to be naturally truncated forms of the N-terminal half of 130-140 kDa Cry proteins. Interestingly some of these natural truncates can apparently be processed more extensively at the N-terminus in vitro than the CryI toxins (Fig. 1). Protease cleavage sites have been identified in vitro at amino acid 159 in CryIIIA (Carroll et af., 1989), 145 in CryIIA (Nicholls et al., 1989), 236 in CryIVA and 204 in CryIVB (Angsuthanasombat et af., 1993; C. Angsuthanasombat, personal communication). Interestingly these cleavage sites are predicted to be in the loop regions between helices a 3 and a 4 (CryIIIA and CryIIA) and a5-a6 (CryIVA and B). Proteins cleaved in vitro at these sites retain activity in vivo, but it remains possible that the N-terminal fragment remains closely associated with the rest of the toxin molecule after nicking.

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It is not known whether this N-terminal nicking occurs in vivo, but if so it might aid insertion of part of the toxin into the target membrane (see Section 5). CryIVD is apparently unique amongst Cry toxins in its mechanism of activation. The 65-kDa protoxin is cleaved into two parts, 30 and 35 kDa (Chilcott and Ellar, 1988). It is not yet known whether these polypeptides remain associated with each other, which is the active moiety, or whether the two parts may act together as a binary toxin. The full-length 27-kDa CytA protein has a low cytolytic action in vitro, but its activity is increased after proteolytic cleavage to a 25-kDa product (Chilcott and Ellar, 1988). It may thus not be a true protoxin (although it is difficult to exclude the possibility that the activity of the 27-kDa protein is due to contamination with small amounts of the 25-kDa activated toxin). In contrast the full-length CytB protein is completely inactive in vitro before proteolytic activation (Knowles et al., 1992). The main difference between the two known Cyt toxins is a C-terminal extension of 15 amino acids in CytB, which may be responsible for an altered activation requirement (Koni and Ellar, 1993). 4.3

THE PERITROPHIC MEMBRANE

Since the foregut of all insects susceptible to Bt toxins is lined with impermeable cuticle, the first “membrane” encountered by ingested toxins is the peritrophic membrane, so it is interesting to note that some Bt toxins bind extensively and apparently with a degree of specificity to this structure (Bravo et al., 1992a, b). Binding was not correlated with toxicity, but in the case of two closely related toxins which both bound to the columnar cell brush border membrane, CryIA(b) bound to the peritrophic membrane but CryIA(c) did not (Bravo et al., 1992a). A major role of the peritrophic membrane is to protect the gut from foreign bodies, and it is tempting to speculate that possession of toxin binding sites may be one line of defence by the insect against Bt attack. The peritrophic membrane prevents the passage of aggregates (but not monomers) of Bt toxins by a sieving effect (Yunovitz et al., 1986), and chitinases act synergistically with Bt, possibly by digesting the peritrophic membrane (Smirnoff, 1974; Sneh et al., 1983).

4.4 4.4.1

RECEPTORS

Cry toxin receptors

After penetrating the peritrophic membrane the lepidopteran toxins of Bt are known to bind with high affinity to specific receptors found on the brush border of the columnar cells lining the midgut. This has been convincingly


289

demonstrated in vitro by kinetic binding and competition studies using CryI toxins on brush border membrane vesicles from lepidopteran larvae (Hofmann et af., 1988a, b; Van Rie et af., 1989, 1990a). These studies have revealed great complexity in binding patterns: a given toxin may have more than one class of binding site in a single insect, and more than one type of toxin may compete for binding to a single site (reviewed by Wolfersberger, 1990a; Gill et af., 1992). Similar studies of the binding of CryIIIA to brush border membrane vesicles from the beetle gut suggested that this toxin bound with lower affinity than Cry1 toxins (Slaney et al., 1992). Recently the site of toxin binding in vivo has been demonstrated directly by visualization of the location of ingested toxins (Bravo et af., 1992b). This study confirms that CryI toxins bind specifically to the brush border membrane of midgut columnar cells in susceptible Lepidoptera, and shows for the first time that the CryIIIA toxin binds to the brush border membrane of the gut epithelial cells of a beetle ( L . decernZineata), in the latter case only in the posterior region of the midgut. The toxins were not detected inside the cells or in the goblet cavity, even several hours after the first cytopathic symptoms could be observed (Bravo et af., 1992b). A similar study on the binding site for mosquitocidal toxins in vivo demonstrated that both the CryIVD and the CytA toxins bound to a specific region of the posterior midgut of mosquito larvae (Ravoahangimalala et af., 1993). Although specific binding to brush border membranes is not always quantitatively correlated with toxicity (Wolfersberger, 1990b; Ferrk et al., 1991; Garczynski et af., 1991) it appears to be an absolute requirement for toxicity. The putative receptors for at least some Bt CryI toxins are glycoproteins of 120-180 kDa (Knowles and Ellar, 1986; Garczynski et af., 1991; Knowles et af., 1991; Oddou et af., 1991). The normal physiological function of these membrane glycoproteins is not known, nor is their role in toxin action, and will probably not be elucidated until a functional receptor is cloned and characterized. Possible functions for the receptor in Bt toxicity are reviewed by Knowles and Dow (1993). At very high doses some Cry toxins are apparently able to dispense with a receptor and insert into artificial phospholipid bilayers, rendering the bilayers permeable to ions or small molecules (Yunovitz and Yawetz, 1988; Haider and Ellar, 1989; Slatin et af., 1990; English et af., 1991). However, addition of brush border membrane proteins from a susceptible insect reduces the required dose of toxin up to 1000 fold (English et af., 1991; Slaney et al., 1992). The specificity of most Bt toxins in vitro to cell lines derived from susceptible insects suggests that, unlike Cyt toxins, Cry toxins at physiological concentration do not insert spontaneously into natural membranes in the absence of high affinity receptors.

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4.4.2 Cyt toxin receptors Unlike Cry toxins, Cyt toxins have a broad-spectrum cytolytic activity in vitro, lysing most eukaryotic cells (Thomas and Ellar, 1983a; Drobniewski and Ellar, 1989; Knowles et al., 1992) and rendering phospholipid bilayers leaky at low toxin concentrations (Drobniewski and Ellar, 1988; Knowles et al., 1989, 1992). The finding that Cyt toxins bind specifically to certain unsaturated phospholipids led to the theory that these toxins do not require a protein receptor, but insert spontaneously into membranes containing these ubiquitous phospholipids (Thomas and Ellar, 1983b; Drobniewski and Ellar, 1989). However, recent results suggest that even for these toxins there may be a specific receptor in vivo: binding of Cyt toxins to lipid membranes and activity against non-target cells in vitro decreases with increasing pH (Maddrell et al., 1989; Knowles et al., 1990a) but high pH is a feature of the midgut of insects killed by Cyt toxins; a mutated CytA toxin with a single amino acid substitution is defective in spontaneous insertion into lipid bilayers but retains specific action against mosquito cells (Ward et al., 1988; Knowles et al., 1990b); CytA and CytB are more toxic to mosquito cells in vitro than to other cells (Chilcott and Ellar, 1988; Knowles et al., 1992); and CytA binds to a specific region of the mosquito midgut in vivo (Ravoahangimalala et a f . , 1993). 4.4.3 Receptors and resistance Despite increasing and long-term use of Bt preparations as commercial insecticides there have been very few reports of resistance in the field. Resistance to Bt toxins and its management are reviewed by McGaughey and Whalon (1992). In two laboratory-selected strains and one field strain the mechanism of resistance involved a decrease in the number or affinity of toxin receptors (Van Rie et al., 1990b; Ferre et al., 1991; MacIntosh et al., 1991). Interestingly, in one study the resistant insects showed an increase in the number of binding sites for a Cry toxin to which they had not previously been exposed (Van Rie et al., 1990b), and in another a decrease in the affinity of the receptors for two Bt toxins was accompanied by an increase in the number of toxin receptors (MacIntosh et al., 1991). These results could be interpreted as evidence that the toxin receptors fulfil an important physiological role in the insect, so that any alteration in the receptor must be accompanied by an increase in numbers of the same or a related protein in order to maintain their normal activity. The recent discovery of a laboratory-selected strain of Heliothis virescens displaying cross-resistance to Bt toxins which apparently do not share a receptor raises urgent concerns about strategies to prevent such resistance occurring in the field. The resistance mechanism is not known, but does not involve an alteration in number or affinity of toxin binding sites in the insect gut (Gould et al., 1992).


4.5

29 1

PORE FORMATION

After binding to a specific receptor on the brush border membrane of insect midgut epithelial cells, Bt toxins insert rapidly and irreversibly into the plasma membrane of the gut cell (Wolfersberger et al., 1986; Van Rie et al., 1989). There is evidence from in vitro studies that (at least in the case of CytA) the proteins then form oligomers of up to 16 toxin monomers (Maddrell et al., 1988; Chow et al., 1989). The next step involves the formation of a pore or lesion in the plasma membrane. It is not yet known whether the toxin alone forms the pore in vivo, or whether the toxin and receptor together form a complex. The selectivity of Cry toxin-induced pores is reviewed by Knowles and Dow (1993). Indirect assays, using the ability of Bt toxins to disrupt cation/ amino acid symport as a measure of the cation permeability of the toxic lesion, suggested that the pore was K+- or cation-selective (Sacchi et af.. 1986; Wolfersberger, 1989). The channels formed in liposomes (English et al., 1991) and planar lipid bilayers (Slatin et af., 1990) by high doses of Cry toxins (see Section 4.4.1) show some selectivity to cations over anions, although in bilayers this is only by a factor of 25:1 (Slatin et al., 1990). A patch clamp study of a cell line treated with CryIC suggested that the toxin was able to act from either side of the plasma membrane to induce a small anion channel (Schwartz et al., 1991). In contrast, direct permeability assays on cell lines (Knowles and Ellar, 1987) and brush border membrane vesicles (Hendrickx et af., 1989; Carroll and Ellar, 1993) supported the notion of a non-selective toxin-induced pore of about 0.6-nm radius, permeable to cations, anions and uncharged molecules up to the size of sucrose. Despite the lack of sequence homology between Cry and Cyt toxins it appears that both classes of toxin form similar lesions in affected cells (Knowles and Ellar, 1987; Drobniewski and Ellar, 1988; Knowles et al., 1989; Knowles et af., 1992). It is possible that some of the reported discrepancies in the characteristics of toxin-induced pores result from differences in the experimental method employed, for instance the CrylC-induced channel was selective for anions at pH 6.2 but became more cation-selective at higher pH (Schwartz et al., 1991). However, it remains possible that there are real toxin- or insect-specific differences in the size or selectivity of toxic lesions.

4.6

CELL LYSIS

Pore formation is a common mechanism of action for cytolytic bacterial protein toxins (English and Slatin, 1992). The outcome of inserting a pore in a plasma membrane will depend on the environment and activity of the cell. In most cells, the resting potential is maintained by a combination of a Na+/

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K+-ATPase and a K+ permease. Selective elevation of the membrane permeability to K+ might thus have little effect on most cells since they are usually already more permeable to K+ than to other ions. However, this condition does not pertain to the columnar cells of the larval lepidopteran midgut. The apical membrane of these cells is impermeable to K + , since the K+ electrochemical gradient is used to drive nutrient uptake (Harvey et al., 1987). The mechanism by which the Bt toxin-induced pore causes the observed damage to insect gut cells is explained by two hypotheses, discussed in Sections 4.6.1 and 4.6.2. The first theory applies to Lepidoptera (and possibly to other insects with a high pH gut), while the second is applicable to all insects affected by Bt toxins. 4.6.1 The “proton peril” hypothesis One method of investigating the mode of action of Bt toxins on the insect gut is to study their effect on an isolated gut under voltage clamp conditions. In the larval lepidopteran gut the current required to maintain a transepithelial potential of zero (the short-circuit current) is largely, though not exclusively, a measure of the activity of the “K+ pump”: the proton pumping V-ATPase linked to a K+/H+ exchanger. Unfortunately most experiments carried out using this method have utilized Bt protoxins (often a mixture of Cry proteins), solubilized at high pH but not proteolytically activated. Such experiments have reported complete or partial inhibition of the short-circuit current within 10-40 min of protoxin application (Harvey and Wolfersberger, 1979; Gupta et al., 1985; Crawford and Harvey, 1988). Using activated toxin complete inhibition occurs in under 5 min (J. A. T. Dow and S. H. P. Maddrell, personal communication). Harvey and Wolfersberger (1979) found that a dose of protoxin which inhibited the short-circuit current by 70% within 40 min had no effect on the flux of K+ from blood to lumen but increased the flux in the opposite direction threefold and decreased transepithelial resistance by 50%. These results supported the proposal that the toxin induced or formed a K+-selective channel in the columnar cell apical membrane, making the columnar cells leaky to K+ but not affecting active K+ pumping in the goblet cell (Harvey et al., 1986; Crawford and Harvey, 1988). Such studies led to the hypothesis that formation of a cation leak in the normally K+-impermeable columnar cell apical membrane would result in depolarization of this membrane and a consequent efflux of H + down the large pH gradient: the apical membrane separates compartments with a greater than 10 000-fold H+ concentration gradient. The rise in columnar cell cytoplasmic pH would kill the cells, leading to gut breakdown and eventual death of the larva (Harvey et al., 1986; Wolfersberger, 1992). Parenthetically, it is interesting to note that the inhibition of the transepithelial short-circuit current by Bt toxins could be reversed in the


293

presence of 4-6 mM Ba2+ or Ca2+ (Crawford and Harvey, 1988). It was proposed that this reversal was due to closure by the divalent cation of the toxin-induced pore (Crawford and Harvey, 1988). However subsequent studies using brush border membrane vesicles showed that Ba2+ and Ca2+ did not have a direct effect on the toxic lesion (Hendrickx et a / . , 1989; Wolfersberger, 1989) and it was suggested that their effect on the intact gut might be to close gap junctions, isolating the damaged cells and thus allowing K+ pumping to continue (Hendrickx et af., 1989). One might alternatively speculate that if isolation of the goblet cells from toxindamaged columnar cells causes basal entry of K + to become the limiting factor in K+ pump activity (as discussed in Section 4.6.2) an effect of Ba2+ on the basal side may be due to its ability to activate K+ channels in the goblet cell basal membrane (Moffett and Lewis, 1990).

4.6.2

The osmotic lysis hypothesis

Cells have a continuous tendency to take in water because of the Donnan effect exerted by the fixed negative charges of intracellular macromolecules such as proteins and nucleic acids. To counter this tendency cells pump small ions into the external medium. A minor leak in the plasma membrane will be counteracted by the activity of ion pumps, but a major breach in the membrane or inhibition of the transport processes inevitably condemns the cell to osmotic lysis. In studies of the effect of Cry and Cyt toxins on insect cell lines in vitro. it was found that addition of osmotic protectants, of the size of the trisaccharide raffinose and above, protected the cells from toxin-induced lysis (Haider and Ellar, 1987; Knowles and Ellar, 1987; Drobniewski and Ellar, 1988). It was proposed that Bt toxins killed cell lines by the process of colloid-osmotic lysis described above. Histopathological studies of insect guts damaged by Bt toxins in vivo show a common process of blebbing, swelling and budding of microvilli followed by cell swelling and lysis (de Barjac, 1978; Endo and Nishiitsutsuji-Uwo, 1980, 1981; Liithy and Ebersold, 1981; Percy and Fast, 1983; Bauer and Pankratz. 1992; Bravo et al., 1992b). Although insects with a steep pH gradient across the midgut apical membrane are likely to suffer pH-mediated damage to gut cells with toxin-induced lesions, osmotic lysis will also occur unless the toxic lesions are large enough to allow immediate leakage of intracellular macromolecules. Knowles and Dow (1993) produced the following model to explain the action of Bt toxins on the lepidopteran gut (Fig. 4). The formation of nonselective pores in the columnar cell apical membrane results in entry of K+ and efflux of H + , which rapidly depolarizes this membrane. Small anions can probably also enter through the toxin pore. Both the rise in intracellular pH and membrane depolarization would probably lead to closure of gap

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junctions (Loewenstein, 1981), isolating the goblet cells from the damaged columnar cells. The columnar cells, containing macromolecules which cannot leak out through a 0.6-nm pore, would absorb water osmotically and thus swell and burst by the process of colloid-osmotic lysis (Knowles and Ellar, 1987). Elevated cytoplasmic pH probably accelerates the death of the columnar cells (Harvey et al., 1986). The goblet cells, which actively pump K+ into the goblet cavity, may rely on a supply of K f mainly via the columnar cells (Dow, 1992). When isolated from the columnar cells, a decreased availability of K+ would lower the activity of the K+/H+ exchanger (Wieczorek, 1992), allowing the V-ATPase to acidify the goblet cavity. This will inevitably lead to inactivation of the V-ATPase, either because the resulting high cytoplasmic pH inhibits mitochondria1 ATP synthesis or because the pump builds up a large apical membrane potential which cannot be dissipated in the absence of K+/H+ exchange. Although the goblet cells also contain osmotically active macromolecules, they may lose water to the mucopolysaccharide matrix filling the goblet cavity once their transport capability is compromised (Dow et al., 1984; Moffett and Koch, 1992). This model may explain the histological observation that shortly after a susceptible insect ingests a Bt toxin the columnar cells swell, and after a lag period the goblet cytoplasm condenses while the goblet cavity swells (Endo and Nishiitsutsuji-Uwo, 1980; Bravo et al., 1992b). The physiology of mosquito and beetle larval guts has not been studied as intensively as that of the lepidopteran gut, mainly because of their small size. It is therefore difficult to predict on theoretical grounds the early effects of insertion of a non-selective pore in their gut cell apical membrane. In those larvae with high pH gut contents it is likely that elevated cytoplasmic pH will hasten the death of gut cells. This will not be the case for the main targets of the CryIIIA toxin, whose gut pH is slightly acidic. In all susceptible insects studied, Bt toxins cause rapid swelling of the target epithelial cells, so it is probable that colloid-osmotic lysis is a common feature in the mechanism of action of Bt toxins, both for gut cells in vivo and for insect cell lines in vitro.

5

Models for the mechanism of pore formation

Chilcott et al. (1990) proposed a model for the orientation of CytA inserted into a plasma membrane, based on structural predictions (Ward et al., 1988) and protease cleavage patterns of the inserted protein (Knowles et al., 1990a). Cry toxins also insert into target membranes in the presence of receptors or (at high concentrations) in the absence of receptors (see Section 4.4.1). There is evidence that the pores formed in planar lipid bilayers in the absence of receptors are different in size and kinetics to those formed in


K+ H+

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H20

FIG. 4 Model for the action of Bt toxins on lepidopteran gut cells. Short-term effects (0-30 min): the toxin generates a non-selective pore in the columnar cell apical membrane. A large influx of K: is driven by the apical membrane potential and a smaller efflux of H + is driven by the p H gradient. Anions may also permeate the toxin-induced lesion. Depolarization of the apical membrane and rise in cytoplasmic p H trigger closure of the gap junctions (1). The columnar cells swell as water enters osmotically. The goblet cells, isolated from the damaged columnar cells. eventually stop active K+ transport as activity of the K+/Hf exchanger decreases (2). Long-term effects (not shown): columnar cells lyse osmotically and the permeability barrier of the gut is breached. The V-ATPase (3) is inhibited. Water moves osmotically from the goblet cells to the goblet cavity matrix and the goblet cells shrink as their cavities swell.

their presence (M. G. Wolfersberger, personal communication) so we should be cautious about assuming that the mechanism of pore formation is the same in each case. Bearing this caveat in mind, it is still possible to suggest mechanisms by which Cry toxins might insert into membranes, with or without the assistance of a receptor. By analogy with other membraneinserting toxins (Lakey et al., 1991; Choe er al., 1992) and with transmembrane ion channels, the most likely structures to penetrate the membrane and form a pore are amphipathic a-helices of at least 20 amino acids or, less commonly, a p-barrel. The structures in direct contact with the hydrophobic core of the plasma membrane should be of sufficient length to

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span the bilayer and must present a hydrophobic face to the membrane lipid. Such structures can be recognized by various computer algorithms, but it should be remembered that the pore might be lined by shorter helices or sheets which need not contain hydrophobic residues and are unlikely to be recognized by existing algorithms (Lodish, 1988). Pores may be formed by a single toxin molecule which spans the bilayer several times, or by oligomers of toxin molecules (Bhakdi and Tranum-Jensen, 1987). Hodgman and Ellar (1990) compared the sequences of Cry toxins from different classes and found no putative fi-strands suitable for formation of transmembrane fi-barrels. They did identify up to six a-helices which showed a striking conservation of amphipathic character despite divergence in amino acid sequence between individual toxin classes. On the basis of these sequence comparisons and structural predictions the authors proposed a general model for pore formation by Cry toxins. In their favoured model a hexameric toxin pore was lined by six helical hairpins, each donated by a toxin molecule. A hexamer was proposed on the grounds of geometrical considerations of the estimated area of the hydrophobic faces and internal pore radius of 0.6 nm (estimated by Knowles and Ellar, 1987), but has no direct experimental support. The X-ray crystal structure of CryIIIA (Li et al., 1991) showed that the Nterminal half of the toxin (domain I) does indeed contain a number of amphipathic a-helices of sufficient length to span a membrane (Fig. 5A). This domain is composed of six amphipathic helices surrounding a central hydrophobic helix (helix as). The hydrophobic surfaces of the outer helices face inwards, the opposite arrangement to that envisaged for an aqueous channel. We would thus expect that a large conformational change must occur if part or all of this domain inserts into the insect plasma membrane. There are a number of possible mechanisms for the insertion of a Cry toxin into its target membrane. Two models are discussed below and shown in Fig. 5. Both models assume that binding of the toxin to its receptor triggers the conformational changes needed for insertion, but that the receptor does not form part of the pore.

5.1

THE “PENKNIFE” MODEL (FIG. S B )

Hodgman and Ellar (1990) nominated helices a5 and a6 (ph 4 and 5 in their nomenclature) as the pair most likely to form the pore. This was on the basis of the predicted area of the hydrophobic faces of the amphipathic helices. Alignment of other Cry sequences with the CryIIIA structure shows that these two helices are highly conserved amongst different Cry toxins (Li et al., 1991). Helices a5 and a6 are joined at the end of domain I predicted to be furthest away from the membrane and would therefore have to flip out of domain I like a penknife opening (Fig. 5B). (A “penknife opening” of a


297

FIG. 5 Models for insertion of Bt toxins into the insect plasma membrane, based on the CryIIIA structure. The a helices in domain I are numbered from the Nterminus, according to Li ef al. (1991). The receptor is indicated by R. (A) One possible orientation of the toxin as it binds to its receptor. A conformational change triggered by binding could be transmitted from domain I1 to domain I via a7 (Li et al., 1991), initiating membrane insertion of two or more helices. The conserved sequence blocks described in Fig. 1 are stippled. (B) The “penknife” model. modified from Hodgman and Ellar (1990): helices a5 and a6 flip into the membrane as a helical hairpin. (C) The “umbrella” model, modified from Li et al. (1991): helices a4 and a5 drop down into the plasma membrane as a helical hairpin, and the other helices flatten out on the membrane surface, their hydrophobic faces towards the membrane. In both models insertion of a pair of helices may be followed or accompanied by formation of an oligomeric barrel-stave pore with a central aqueous channel.

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helical hairpin was proposed to be an early step in the association of colicin A with a membrane surface prior to insertion (Lakey et al., 1992).) This model does not require rearrangement of the rest of domain I, although a4 would probably have to slide downwards relative to a3. An aqueous pore could be formed by oligomerization of a number of toxin molecules.

5.2

THE “UMBRELLA” MODEL (FIG. 5c)

Li et al. (1991) favoured a model in which pairs of helices a6 and a7, or a 4 and a5, insert as a helical hairpin and initiate the rearrangement of the rest of domain I to form a pore. These helix pairs are joined at the side of domain I expected to be closest to the plasma membrane, and thus could drop down into the membrane while the remaining helices opened on to the membrane surface like the ribs of an umbrella (Fig. 5C), a mechanism proposed for colicin A (Lakey et al., 1991). There may need to be proteolytic nicking at one or more interhelical loops to permit this large conformational change (Li et al., 1991) (see Section 4.2). The pore may be formed by oligomerization or by further insertion of other helices from domain I. Both of these models are proposed on theoretical grounds by comparison with membrane channels or channel forming toxins. Initiation of the insertion event might require a conformational change induced by receptor binding; might occur spontaneously when the toxin is close enough to the hydrophobic membrane; might require proteolysis of one or more interhelical loops or might follow oligomerization of toxin molecules. The actual mechanism may be revealed by identifying which parts of the toxin insert into the membrane, and ultimately by solution of the crystal structure of the toxin-induced pore.

6 Conclusions and future prospects

There are still many unanswered questions about the mechanism of action of Bt &endotoxins. Most of the research has focused on the Cry1 toxins and their target insects. Paradoxically, CryIIIA, the only Bt toxin whose threedimensional structure has been solved, is one about which we know very little in other respects, since the insects it kills are very small, and its insolubility at neutral pH makes it difficult to assay in vitro. The crystal structure of CryIIIA now permits site-directed mutagenesis and segment swapping experiments to be designed in a rational manner in order to identify the functional domains of the toxins. One attractive prospect might be the replacement of the specificity domain with a binding domain directed


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to a target of choice, opening up the possibility of “designer” pesticides or a new class of immunotoxins. Characterization of toxin receptors and investigation of the mechanism of toxin synergism may assist in strategies for resistance management, a vital concern if Bt is to maintain its important position as the most extensively used biological insecticide.

Acknowledgements

I am grateful to Chanan Angsuthanasombat, Chris Chilcott, Joe Carroll, Julian Dow, Pandelakis Koni and Mike Wolfersberger for allowing me to include their results prior to publication, and David Ellar for lively debates on the mechanism of toxin action and insertion, and for providing the “umbrella model”. I thank Julian Dow, David Ellar and Emmy Van Kerkhove for critical comments on the manuscript, and John Rodford and Jade Li for assistance in preparation of the figures. I thank The Royal Society for financial support of my own unpublished research mentioned in this review.

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thuringiensis subsp. kurstaki &endotoxin inhibition of potassium current across isolated rnidgut of larval Manduca sexta. J. exp. Biol. 137, 277-286. Crowson, R. A. (1981). “The Biology of the Coleoptera”. Academic Press, London. Dadd, R. H. (1975). Alkalinity within the midgut of mosquito larvae with alkalineactive digestive enzymes. J. Insect Physiol. 21, 1847-1853. Dadd, R. H. (1976). Loss of midgut alkalinity in chilled or narcotized mosquito larvae. Ann. Entomol. SOC.Am. 69, 248-254. Dalhammar, G. and Steiner, H. (1984). Characterization of inhibitor A. a protease from Baciflus thuringiensis which degrades attacins and cecropins, two classes of antibacterial proteins in insects. Eur. J. Biochem. 139, 247-252. de Barjac, H. (1978). Etude cytologique de I’action de Bacillus thuringiensis var. israelensis sur larvae de moustiques. C.R. Acad. Sci. Paris ser. D 286, 1629-1632. Dow, J. A. T. (1984). Extremely high pH in biological systems: a model for carbonate transport. A m . J. Physiol. 246, R633-R635. Dow, J . A . T. (1986). Insect midgut function. Adv. Insect. Physiol. 19, 187-238. Dow, J. A. T. (1992). pH gradients in lepidopteran midgut. J. exp. Biol. 172, 355-375. Dow, J . A. T. and Peacock, J . M. (1989). Microelectrode evidence for the electrical isolation of goblet cell cavities in Manduca sexta middle midgut. J . exp. Biol. 143, 101-114. Dow, J. A. T., Gupta, B. J . , Hall, T. A. and Harvey, W. R. (1984). X-ray microanalysis of elements in frozen hydrated sections of an electrogenic K+ transport system: the posterior rnidgut of tobacco hornworm (Manduca sexta) in vivo and in vitro. J. Membr. Biol. 77, 223-241. Drobniewski, F. A. and Ellar, D. J. (1988). Investigation of the membrane lesion induced in vitro by two mosquitocidal &endotoxins of Bacillus thuringiensis. Curr. Microbiol. 16, 195-199. Drobniewski, F. A. and Ellar, D. J . (1989). Purification and properties of a 28kilodalton hemolytic and mosquitocidal protein toxin of Bacillus thuringiensis subsp. darmstadiensis 73-E10-2. J . Bacteriol. 171, 306C3067. Dulmage, H. T. and Aizawa, K . (1982). Distribution of Bacillus thuringiensis in nature. In “Microbial and Viral Pesticides” (Ed. E. Kurstak), pp. 209-237. Marcel Dekker, New York. Earp, D. J . and Ellar, D. J. (1987). Bacillus thuringiensis var. morrisoni strain PG14: nucleotide sequence for a gene encoding a 27-kDa crystal protein. Nucleic Acids Res. 15, 3619. Ebersold, H. R., Luthy, P., Geiser, P. and Ettlinger, L. (1978). The action of the 6endotoxin of Bacillus thuringiensis: an electron microscope study. Experientia 34, 1672. Ellar, D. J . (1990). Pathogenicity determinants of entomopathogenic bacteria. In “5th International Colloquium on Invertebrate Pathology and Microbial Control” (Ed. D. Pinnock), pp. 298-302. Society for Invertebrate Pathology, Adelaide, Australia. Endo, Y. and Nishiitsutsuji-Uwo, J. (1980). Mode of action of Bacillus thuringiensis b-endotoxin: histopathological changes in the silkworm midgut. J. lnvertebr. Pathol. 36, 90-103. Endo, Y. and Nishiitsutsuji-Uwo, J. (1981). Mode of action of Bacillus thuringiensis 8-endotoxin: ultrastructural changes of midgut epithelium of Pieris, Lymanrria and Ephestia larvae. Appl. Ent. Zool. 16, 231-241. English, L. and Slatin, S. L. (1992). Mode of action of delta-endotoxins from Bacillus thuringiensis: a comparison with other bacterial toxins. Insect Biochem. Mol. Biol. 22, 1-7.

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Harvey, W. R., Cioffi, M., Dow, J . A. T. and Wolfersberger, M. G. (1983). Potassium ion transport ATPase in insect epithelia. J . exp. Biol. 106, 91-117. Harvey, W. R., Cioffi, M. and Wolfersberger, M. G. (1986). Transport physiology of lepidopteran midgut in relation to the action of B.t. delta-endotoxin. In “Fundamental and Applied Aspects of Invertebrate Pathology” (Eds J . M. Vlak. D. Peters and R. A. Samson), pp. 11-14. Grafisch bedrijf Ponsen and Looijen, Wageningen, The Netherlands. Harvey, W. R., Crawford, D. N., Eisen, N. S., Fernandes, V. F., Spaeth, D. D. and Wolfersberger, M. G. (1987). The potassium impermeable apical membrane of insect epithelia: a target for the development of safe insecticides. Mem. Inst. Oswaldo Cruz 82, 29-34. Heimpel, A. M. and Angus, T. A. (1959). The site of action of crystalliferous bacteria in Lepidopteran larvae. J . Insect Pathol. 1, 152-170. Heimpel, A. M. and Angus, T. A. (1960). Bacterial insecticides. Bacteriol. Rev. 24, 266288. Held, G. A , , Kawanishi, C. Y. and Huang, Y.-S. (1990). Characterization of the parasporal inclusion of Bacillus thuringiensis subsp. kyushuensis. J . Bacteriol. 172, 481-483. Hendrickx, K., de Loof, A. and Van Mellaert, H . (1989). Effects of Bacillus thuringiensis delta-endotoxin on the permeability of brush border membrane vesicles from tobacco hornworm (Manduca sexta) midgut. Comp Biochem. Physiol. 95C, 241-245. Hodgman, T. C. and Ellar, D. J. (1990). Models for the structure and function of the Bacillus thuringiensis S-endotoxins determined by compilational analysis. D N A Sequence 1, 97-106. Hofmann, C., Liithy, P., Hutter, R. and Pliska, V. (1988a). Binding of the deltaendotoxin from Bacillus thuringiensis to brush-border membrane vesicles of the cabbage butterfly (Pieris brassicae). Eur. J . Biochem. 173, 85-91. Hofmann, C., Vanderbruggen, H., Hofte, H., Van Rie, J., Jansens, S. and Van Mellaert, H . (1988b). Specificity of Bacillus thuringiensis &endotoxins is correlated with the presence of high-affinity binding sites in the brush border membrane of target insect midguts. Proc. Natl. Acad. Sci. USA 85, 78447848. Hofte, H. and Whiteley, H. R. (1989). Insecticidal crystal proteins of Bacillus thuringiensis. Microbiol. Rev. 53, 242-255. Tshiwata, S. (1901). On a kind of severe flacherie. Dainihon Sanshi Kaiho 9, 1-5 (in Japanese). Keil, C. B. (1991). Field and laboratory evaluation of a Bacillus thuringiensis var. israelensis formulation for control of fly pests of mushrooms. J . econ. Entomol. 84, 1180-1188. Knowles, B. H. and Ellar, D. J. (1986). Characterisation and partial purification of a plasma membrane receptor for Bacitlus thuringiensis var. kurstaki lepidopteranspecific S-endotoxin. J . Cell Sci. 84, 89-101. Knowles, B. H. and Ellar, D. J. (1987). Colloid-osmotic lysis is a general feature of the mechanism of action of Bacillus thuringiensis S-endotoxins with different insect specificity. Biochim. Biophys. Acta 924, 509-518. Knowles, B. H., Blatt, M. R., Tester, M., Horsnell, J. M., Carroll, J., Menestrina, G. and Ellar, D. J. (1989). A cytolytic S-endotoxin from Bacillus thuringiensis var. isruelensis forms cation-selective channels in planar lipid bilayers. FEBS Lett. 244, 259-262. Knowles, B . H., Carroll, J., Horsnell, J. M. and Ellar, D. J. (1990a). Interactions of a cytolytic toxin from Bacillus thuringiensis var. israelensis with liposomes and membranes. Zentralblatt Bakteriologie, Suppl. 19, 207-208.

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Yunovitz, H., Sneh, B., Schuster, S . , Oron, U., Broza, M . and Yawetz, A . (1986). A new sensitive method for determining the toxicity of a highly purified fraction from b-endotoxin produced by Bacillus thuringiensis var. entomoocidus on isolated larval midgut of Spodoptera littoralis (Lepidoptera, Noctuidae). J . Invertebr. Pathol. 48, 223-231.

Insect Glutamate Receptors P. N. R. Usherwood Department of Life Science, University of Nottingham, Nottingham NG7 2RD, UK

Introduction 309 Ionotropic glutamate receptors of insect central nervous systems 310 Ionotropic glutamate receptors of skeletal muscle 314 3.1 Structure - activity studies 315 3.2 Channel gating kinetics of extrajunctional glutamate receptors 316 3.3 Developmental studies of skeletal glutamate receptors 329 3.4 Ibotenate-sensitive glutamate receptors of skeletal muscle extrajunctional membrane 329 Glutamate receptors of visceral muscle 330 Metabotropic glutamate receptors 331 Cloning, expression and functional characterization of glutamate receptors 332 6.1 Exogenous expression of native RNAs 332 6.2 Cloning and structural characterization of central and peripheral glutamate receptors 333 Summary 334 Acknowledgements 334 References 334

1 Introduction

Over 20 years have past since I last wrote on insect glutamate receptors (GluR) for Advances in Insect Physiology (Usherwood, 1968). At that time, the idea that receptors such as these might occur at synapses in arthropods was generally accepted, but proposals that they might also mediate transmission at synapses in vertebrate central nervous systems (CNS) were still a matter of controversy. The ubiquitous distribution of L-glutamic acid in excitable and other tissues, and its supposedly high intracellular and, sometimes, extracellular concentrations were the major arguments raised against its proposed role as a vertebrate neurotransmitter (reviewed by Usherwood, 1976, 1978). The present position is remarkably different. Glutamate receptors (GluR) of vertebrate CNS have been cloned (Sommer and Seeburg, 1992) and these membrane macromolecules have been implicated in important functions, such as information storage (Collingridge

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and Bliss, 1987), and in a number of disorders of human brain, including cerebral ischaemia (Zivin and Choi, 1991). Before molecular biology took such a firm hold on the field, studies of arthropod excitatory nerve-muscle junctions provided much of the basic information on the properties of CluR which underpins the current huge interest in these proteins, particularly in mammals. Advances made in understanding mammalian GluR following the introduction of molecular biology into neuroscience have been revolutionary, and have temporarily limited the contributions that insect models can make to basic studies in this field. Also, in many areas, our knowledge of GluR in insects now lags behind that of mammalian GluR. A number of excellent reviews of insect GluR have been published (e.g. Leake and Walker, 1980; Piek, 1985; Duce, 1988), and it would not be profitable to cover the same ground again. Instead, I will concentrate on recent developments, whilst making some hopefully pertinent and timely comments on the various relationships between insect GluR and their counterparts in vertebrates. I will end my review with some opinions on possible future developments, and point out how insect nerve and muscle preparations might serve as models of GluR function in normal and diseased human brain. Figure 1 presents a schematic summary of the distribution and types of GluR in insects and is a guide to what follows.

2

lonotropic glutamate receptors of insect central nervous systems

“Insect central nervous systems have provided a major source of inspiration and a fair share of frustration for the industrial scientist wishing to exploit this tissue as a target for novel pesticides” (Usherwood et al., 1980). This comment, made at the first Neurotox ‘79 (1979) symposium of the UK Society for Chemical Industry, remains true to this day. Since that meeting, considerable progress has been made in understanding the properties of cholinergic synapses and acetylcholine receptors of insect CNS, but what progress has been made concerning insect GluR? Well, it is now accepted that glutamatergic synapses are present in insect CNS, although their functional properties are still not well characterized. For example, many of the motorneurones that innervate locust (Locusta migratoria) skeletal musculature also make synaptic connections with other neurones in the CNS of this insect. Since the motorneurones release L-glutamate at their neuromuscular synapses, it seems reasonable to assume that their central synaptic contacts are also glutamatergic (Sombati and Hoyle, 1984). Support for this assumption has been provided by the immuno-histochemical studies of Bicker et al. (1988) and Watson (1988), in which L-glutamate antiserum was applied to sections of honey bee and locust ( L . migratoria) CNS. The

INSECT GLUTAMATE RECEPTORS

31 1

n I CNS

mGluR n

J I

qGluR

iGluR

MUSCLE FIG. 1 Schematic representation of the distribution and types of glutamate receptor (GluR) present in insect central nervous system (CNS) and on skeletal and visceral muscle (MUSCLE). Ionotropic GluR are of two major physiological types: those that gate cation-selective channels, that is, the excitatory receptors or D-receptors (V); and those that gate anion-selective channels, i.e. the inhibitory receptors or H-receptors (V).Both types are often represented on extrajunctional membrane of insect motorneurones. Central glutamatergic synapses involving motorneurones may be either inhibitory or excitatory. At excitatory nerve-muscle junctions of locust (S. gregaria) skeletal muscle there are pharmacologically distinct subpopulations of GluR (1-3), which are quisqualate-sensitive, ibotenate-sensitive and aspartatesensitive respectively. There are three populations of extrajunctional GluR: qGluR, which gate cation-selective channels and which are identical pharmacologically to the quisqualate-sensitive, postjunctional GluR; iGluR, which gate chloride channels and which are sensitive to ibotenate; and mGluR (insect metabotropic receptors), which induce membrane potential changes via secondary messengers. Presynaptic GluR, which are sensitive to NMDA and gate cation-selective channels have been discovered at excitatory nerve-muscle junctions (A). However, they may co-exist with other pharmacological types of o-receptors, with H-receptors and with mGluR at this site. See text for references.

presence of immunoreactivity was demonstrated not only in motorneurones but also in certain interneurone populations. The notion that L-glutamate is a central transmitter in insects received further support when this amino acid was ionotophoresed onto regions in the locust (L. rnigrutoria) metathoracic ganglion where motorneurone projections are located and the nerve cells with which these motorneurones might be expected to synapse were depolarized (Sombati and Hoyle, 1984; Dubas, 1991).

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It was Usherwood et al. (1980) and Giles and Usherwood (1985) who had first discovered GluR in locust (Schistocerca gregaria) CNS, whilst studying the pharmacology of neuronal somata isolated from thoracic ganglia of adults and of cultured larval neurones, but these receptors were clearly extrajunctional (Fig. 1). They responded to L-glutamate, L-aspartate, DLibotenate and L-kainate with either hyperpolarizations or depolarizations or with biphasic changes in membrane potential. Hyperpolarizing responses were obtained by Wafford and Sattelle (1986, 1989) during application of Lglutamate and L-aspartate to an identified in situ neurone (the fast coxal depressor motorneurone, Df ) of cockroach (Periplaneta americana) CNS. In contrast, L-quisqualate, L-kainate and L-cysteate depolarized this cell, whereas N-methyl+-asparatate (NMDA), L-homocysteate and a-amino-3hydroxyl-5-methyl-4-isooxazolepropionic acid (AMPA) were inactive. In further efforts pharmacologically to characterize the Df neurone, Wafford and Sattelle (1989) tested a variety of putative antagonists of vertebrate GluR against the glutamate responses of this cell. Unfortunately, these compounds were either inactive or only weakly active, so it seems possible that GluR of the Df neurone do not have much in common with vertebrate CNS GluR. Dubas (1991) searched for glutamatergic synapses in locust ( L . migratoria) CNS by applying L-glutamate and L-aspartate to dendritic arborizations of flight motorneurones of this insect. These amino acids elicited either a depolarization, a hyperpolarization or a biphasic response, all three responses being accompanied by increases in membrane conductance. It was suggested, on the basis of ion substitution experiments, that both the depolarizing and the hyperpolarizing responses involved chloride activation, but no explanation was given for the biphasic responses. Possibly, some glutamatergic synapses in the locust CNS are inhibitory, although others are clearly not (Sombati and Hoyle, 1984; Burrows et al., 1989). A motorneurone making excitatory glutamatergic synapses with skeletal muscle could, in principle, also make inhibitory and/or excitatory glutamatergic synapses with other excitable cells (Fig. 1). Unfortunately, in most of the above in situ studies, it was not possible unequivocally to determine the exact site of application of GluR agonist, so we do not know whether the actions of these compounds were limited to junctional GluR or whether they also activated extrajunctional GluR. The responses of insect CNS neurones to kainate are of particular interest, because Ultsch et al. (1992) have cloned a kainate-selective subunit from Drosophila melanogaster which is expressed in the CNS of this insect. According to Wafford et a f . (1992), kainate receptors (KainR) of cockroach Df neurone, which are sensitive also to L-quisqualate (ECSo = 160 WM) and L-domoate (ECsn = 7 VM) (ECSofor L-kainate = 263 VM), are pharmacologically distinct from ionotropic KainR of vertebrate CNS, for example they (CNQX), a potent are insensitive to 6-cyano-7-nitroquinoxaline-2,3-dione antagonist of vertebrate KainR. However, their claim that the lack of


313

sensitivity of the Df neurone to AMPA distinguishes cockroach KainR from those of vertebrates is now clearly untenable, because some KainR subunits cloned from mammalian CNS are also insensitive to AMPA (Egebjerg et al., 1991). The outcomes of studies of vertebrate kainate-sensitive GluR suggest that one should be particularly cautious when interpreting pharmacological data for this receptor class because agonists of KainR are not similar with respect to their capacities to desensitize these receptors (e.g. Lambolez ef al., 1990). It seems likely that insect CNS contains more than one type of KainR, because the KainR subunit cloned from D. melanogaster, when expressed as a homo-oligomer in Xenopus oocytes, also fails to respond to AMPA, but its responses to kainate are blocked by CNQX. Until more substantial pharmacological and molecular biological studies are undertaken on GluR of insect CNS the picture concerning the numbers, distributions and subtypes of GluR in this tissue will remain unclear. However, it is not entirely obscure. Some qualitative properties of GluR in the CNS of a few insects have been established. For example, there seems little doubt that somata1 membrane of many neurones in cockroach and locust CNS contain one or two populations of GluR, that is comprising depolarizing receptors (D-receptors), which gate cationic channels and hyperpolarizing receptors (H-receptors), which gate chloride channels. In fact many of the pharmacological and physiological properties of these neuronal receptors are reminiscent of GluR which are present on extrajunctional membrane of locust (S. gregaria) skeletal muscle (Lea and Usherwood, 1973; Cull-Candy and Usherwood, 1973) and adult (not larval and pupal) mealworm (Tenebrio rnolitor) muscle (Saito and Kawai, 1985). A major difference between GluR of CNS and skeletal muscle may be representation in the former of KainR, which do not seem to occur peripherally in insects (Daoud and Usherwood, 1975). It seems reasonable to expect that a better understanding of the types of GluR present in insect CNS neurones will arise from patch clamp studies of these cells, although few such studies have been undertaken so far. Openings of cation-selective channels (50 pS conductance) have been seen in cell-attached patches of cultured neurones of P. americana, made using pipettes containing L-glutamate (Horseman et al., 1988). The channel currents reversed at 0 mV membrane potential. In whole-cell recordings from these cultures, inward currents were recorded at resting potential (c. -30 mV), using patch pipettes containing L-glutamate ( lo p 5 ~ - 1 0 -M)~ dissolved in high sodium/low potassium saline, which reversed at about 0 mV. Interestingly, the frequency of occurrence (fo) of the 50 pS channel openings was increased when concanavalin A was applied to the cells, presumably because the lectin inhibits desensitization of the GluR (see later). There have been few biochemical studies of [3H]glutamate receptor binding sites in insect CNS. Sherby et al. (1987) used a filtration assay to

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study such sites in housefly (Musca domesticus) and honeybee (Apis mellifera) heads and thoracies, and in cockroach (P. americana) nerve cords. In all three cases binding was saturable and stereospecific. KD values ranged from 0.16 to 1.36 p ~ The . concentrations of binding sites were 5.7, 3.8 and 5 pmoVmg protein for housefly brain, honey bee brain and cockroach nerve cord, respectively. These concentrations are lower by two-fold to five-fold than those for insect muscle (Filbin et al., 1985; Sherby et al., 1987), and eight-fold to 18-fold lower than those for rat brain (Foster et al., 1981; Michaelis et al., 1984). The putative receptors of insect CNS and muscle share common pharmacological characteristics, that is, similar binding affinities for L-aspartate, L-quisqualate, L-cysteate and DL-ibotenate (Sherby et al., 1987). L-Kainate and NMDA are either inactive or only weakly inhibit [3H]glutamate binding. However, when interpreting these data it must be remembered that the muscle preparations were probably contaminated with nervous tissue, for example motor nerve terminals which are known to contain GluR (Usherwood and Machili, 1966; Usherwood, 1984), It is surprising, perhaps, that despite the passing of 14 years since that meeting of Neurotox in 1979, progress made in identifying GluR and glutamatergic synapses in insect CNS and in understanding their properties remains rather modest. Added to this is the depressing fact that there have been few, if any, accounts of GluR in insects other than the chosen four, that is, fruitfly, housefly (blowfly), locust and cockroach.

3 lonotropic glutamate receptors of skeletal muscle

GluR are present junctionally and extrajunctionally on locust (S. gregaria and L. migratoria) and mealworm (T. molitor) skeletal muscle fibres, but not as pharmacologically homogeneous populations at either location. This heterogeneity may not be true for other insects, but I would be surprised if it were not. Duce (1988) comprehensively reviewed the contribution of biochemistry to our understanding of GluR of insect skeletal muscle, and pointed out that the results of binding studies on insect muscle membranes have been inconsistent. However, a recent contribution, from Sepulveda and Sattelle (1991), describing a high affinity, saturable ~ - [ ~ H ] g lu ta m abinding te site in a cockroach (P. americana) coxal muscle preparation with a KD of 1.1 p~ and a B,, of 15.5 p m o l h g protein is more promising. It is of particular note on two scores. Firstly, because the pharmacological profile of the L[3H]glutamate binding site corresponds closely with that expected from previous electrophysiological studies and, secondly, because of the discovery of quisqualate-sensitive and ibotenate-sensitive subpopulations of GluR,


31 5

which seem to correspond to the D-receptors and H-receptors, respectively, described electrophysiologically for other insect skeletal muscles.

3.1

STRUCTURE-ACTIVITY STUDIES

L-Quisqualic acid is a potent agonist of a major population of ionotropic GluR of insect skeletal muscle. Quisqualate-sensitive GluR (qGluR) are present postjunctionally at excitatory nerve-muscle junctions in all insects studied so far and extrajunctionally, at much lower population density, in some of these insects. The extrajunctional qGluR are the D-receptors referred to earlier. Single channel studies of extrajunctional qGluR of locust (S. gregaria) muscle have confirmed the high potency of L-quisqualate, which makes this compound a useful tool for studying qGluR. Difficulties of isolating the compound from its natural source (the seeds of Quisqualis indica) have restricted its usefulness. However, a chemical synthesis is now available for both L-quisqualate and its stereoisomer (Bycroft et al., 1984). During some of the first pharmacological investigations of locust skeletal muscle, it was established that D-glutamic acid is only very weakly active on postjunctional GluR, that is, about 1000 times less potent that L-glutamic acid. In other words, these receptors appear to have a high agonist stereospecificity (Usherwood and Machili, 1968; Clements and May, 1974). It came as some surprise, therefore, when the potency of D-quisqualic acid was found to be only c . 8 times less than that of L-quisqualic acid (Boden et al., 1986). In the crystal lattices of these stereoisomers, the nitrogen atom which joins the heterocyclic system to the amino acid residue has a pyramidal configuration, which is comparable to that of the tetrahedron carbon atom in the corresponding position in L-glutamic acid. Therefore, in an aqueous environment the two stereoisomers should be able readily to flip between their two conformers. Computer modelling suggests that only one of the conformers of L-quisqualic acid has the correct shape for binding to qGluR, and that this is also true for D-quisqualic acid. It follows, therefore, that the relative probabilities of binding of these enantiomorphs to qGluR will depend upon their relative abilities to adopt the appropriate binding conformations. An alternative hypothesis is that the ring junction nitrogen of the heterocyclic system of quisqualic acid adopts a trigonal configuration in solution, and that it can readily change from this to a pyramidal geometry (Bycroft and Jackson, 1988). Comparisons of the pharmacological properties of qGluR and extrajunctional H-receptors (ibotenate-sensitive GluR or iGluR) of locust (S. gregaria) muscle, led Lea and Usherwood (1973) to propose that Lglutamate binds to qGluR in a partially-folded conformation and to iGluR in an extended conformation. This idea was tested by Usherwood (1986) using

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single channel analysis to compare the potencies of 4-alkyl-substituted glutamates. Of the four chiral 4-methyl-glutamic acids only the 2S, 4R isomer was active. This established a configurational constraint with respect to the 4-position, that is, analogous to the pyramidal ring joining nitrogen in quisqualic acid (Bycroft and Jackson, 1988). This confirmed that the agonist activity of D-quisqualic acid can be ascribed to the ability of the molecule to undergo a configurational inversion and to employ the full extent of the delocalized anion of the heterocyclic system to assist binding. The configuration inversion hypothesis was tested by comparing the mean open time (mo) and fo for qGluR channels gated by L-quisqualic acid and Dquisqualic acid. Boden et al. (1986) had already found that the relative potencies of D- and L-quisqualate for qGluR of locust muscle are similar to those for excitatory postjunctional GluR. It was discovered that for similar concentrations of these compounds mo was the same for L- and D-quisqualic acid but fo was much higher for L-quisqualic acid (Boden et al., 1986; Kerry and Usherwood, 1993). In other words, the affinities of L- and D-quisqualic acid are similar, but their efficacies are different. These conclusions are consistent with the configuration inversion hypothesis. One hundred and twenty analogues of L-quisqualic acid have been tested at glutamatergic neuromuscular junctions in larval mealworm (T. mofitor) by Miyamoto et al. (1985). The apparent KD for L-glutamate was 1.5 x lop4 M, whereas the equivalent value for L-quisqualate was about 10 times lower. These results are in keeping with previous observations on locust muscle (Clements and May, 1974). Disappointingly, none of the quisqualic acid analogues was even as potent as L-glutamate.

3.2

CHANNEL GATING KINETICS OF EXTRAJUNCTIONAL GLUTAMATE RECEPTORS

Although extrajunctional qGluR and iGluR were discovered almost 20 years ago their functions remain enigmatic. Recent studies of these receptors have related mainly to single channel experiments on metathoracic extensor tibiae muscle fibres of adult locust (S. gregaria). 3.2.1 Quisqualate-sensitive glutamate receptors Channel openings gated by extrajunctional qGluR of adult locust skeletal muscle are readily resolved, because of their high conductance (c. 150 pS) and low population density. The qGluR channel appears to be nonrectifying. Because desensitization of qGluR can be inhibited by concanavalin A (Mathers and Usherwood, 1976, 1978; Evans and Usherwood, 1985), it has been possible to investigate the channel gating kinetics of these receptors in equilibrium studies undertaken over a 10 000-fold range of agonist concentrations. Not surprisingly, single channel studies of qGluR


317

have been in the forefront of developments in the gating kinetics of transmitter receptors. For example, studies of qGluR equilibrated with agonist provided the first indication that the opening rate of a transmitter receptor channel is related to agonist potency (Gration et al., 1981b), the first evidence for influence of agonist concentration on channel open time (Gration et al., 1981a) and the first account of channel state switching (Patlak et al., 1979), which, over a decade later, is just now being described for NMDA receptors of mammalian CNS (Gibb and Colquhoun, 1992). 3.2.1.1 Single channel studies of quisqualate-sensitive glutamate receptors in equilibrium with agonist The equilibrium kinetics of the extrajunctional qGluR of locust muscle have been extensively studied using either megaohm seal, cell-attached membrane patches or gigaohm seal, outside-out patches. In the former case the qGluR were equilibrated with agonist which was present in the patch pipette; in the latter case the agonist was present in the bath saline. In both types of study, membranes were exposed to concanavalin A before agonist application, in order to block desensitization of qGluR. The megaohm seal approach has an advantage over the gigaohm seal technique in that it can be applied to untreated muscle preparations; for gigaohm seal formation it is necessary thoroughly to clean the surface of the muscle fibres with enzyme (e.g. collagenase; Huddie et al., 1986). Initially. only a single concentration (lop4 M) of one agonist (L-glutamate) was used in the equilibrium studies (Patlak et al., 1979; Gration et al., 1982; Kerry et al., 1986, 1987; Kits and Usherwood, 1988). The presence of multiple open and closed states and the state switching, which was so evident in these early studies, led Gration et al. (1982) to the conclusion that the gating kinetics of qGluR are very different from those of nicotinic acetylcholine receptors (nAChR), for example on frog skeletal muscle (Leibowitz and Dionne. 1984). This conclusion was not accepted universally. Cull-Candy et al. (1981) also reported on the gating kinetics of qGluR of locust leg muscle, which they had studied under agonist equilibrium conditions using megaohm seals. Although they observed the state switching phenomenon documented earlier by Patlak et al. (1979), they, nevertheless, proposed that the gating kinetics of the qGluR channel are qualitatively similar to those of the nAChR channel, that is, with a single, biliganded, open state. They gave no explanation for the state switching observed in their study. Later, more detailed and rigorous analyses of qGluR channel gating in cell-attached patches revealed at least three open states and four closed states, and autocorrelation function analysis showed there to be at least three isomerization pathways linking the open states with the closed states (Kerry et al., 1986). These results led to consideration of allosteric schemes for gating of the qGluR channel, that is, as predicted earlier by Gration et at. (1981a, 1982). More recently, Kerry et al. (1988a) used the megaohm seal technique to investigate the influence of L-glutamate concentration on

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the gating kinetics of qGluR, a comprehensive study in which a 10 000-fold concentration range of this agonist was employed. The results of this exercise led to Model 1 which includes four agonist binding sites: 3aKn

4aK"

2aK"

a&"

C dCA U CA2 W CA3 / CA4 GJKB

h ,/aL

1

hI

2K"/KB

1

h2/a2L h2 3aKn

3Knk

l,

h3/u3L h3 2aG

4K"/KB

1

h4/a4L h4

(1)

aCn

OA2V OA3A -4

OA 2k3KBa

3k:"/K,a

4k&/K,a

where: A is an agonist molecule; C and 0 are, respectively, the closed and open states of the receptor channel; KB is the agonist binding constant = 2 7 0 / ~k;:n is the agonist binding rate to the closed receptor-channel = 3 . 7 / ~ / ms; k& is the agonist binding rate to the open receptor-channel = 126/dms; hl-h4 are the channel opening rates = 3.2 X 10-2/ms, 0.69/ms, 3.9Ims and 6.4 x 1OP3/ms respectively; L is the open-closed equilibrium constant for the unliganded receptor-channel complex = 1.5 x and u is the ratio of the agonist binding constant for the open channel to that for the closed channel = 14.2. Model 1 gives a microscopic association constant for the closed channel binding sites of 2.7 X lO'/mol, and a 14.2-fold higher value for the open channel sites. This 10-state Markovian model has been tested for goodness-of-fit i n . comparison with other types (i.e. non-Markovian) of models of channel gating, that is, fractal, diffusion, etc., and has been shown to provide the best description of the observed open and closed time distributions for qGluR (Sansom et al., 1989). One feature of Model 1 is the prediction of cross-correlations between open times and between closed times. Application of cross-correlation function analysis to single channel data for qGluR (obtained in both megaohm seal and gigaohm seal experiments) has provided strong evidence for multiple gateway states and time reversibility of gating of the qGluR channel (Ball et al., 1988). In any analysis of the gating kinetics of an ion channel it is necessary to consider the effect of event omission on the outcome. Openings and closings of brief duration (> 100 ps) are difficult to resolve with current recording procedures and may be lost altogether. The effect of event omission on the behaviour of Model 1 has been considered by Ball et al. (1988), and shown not to influence qualitatively the cross-correlation functions implicit in the model from the relationship between channel open probability (Po) and concentration of g glut am ate. How d o these results and their interpretation compare with those for the qGluR in outside-out patches? Bates et af. (1990) obtained gigaohm


319

recordings from qGluR in outside-out patches excised from collagenasetreated fibres of adult locust (S. gregaria) muscle. Again, analysis of channel open and closed time distributions revealed at least four closed states, whereas dwell-time autocorrelation analysis showed that there are at least three pathways linking the closed states with the open states. A maximum likelihood procedure was used to fit different gating models to the data, and of these models, Model 1 provides the best fit and predicts well most features of the gating kinetics of qGluR. Interestingly, the agonist affinity of qGluR in outside-out patches (i.e. the microscopic association constant (2.7 x 102/mol) for binding of agonist to closed channel sites) is c. 10 times lower than that for qGluR in cell-attached patches. 3.2.2 Agonist Concentration jump studies The routine use of concanavalin A in the equilibrium studies of qGluR is essential. If membrane patches are not pretreated with this lectin and Lglutamate is applied continuously to such patches, openings of qGluR are rarely seen because of desensitization (Gration et a f . , 1980a, b). However, when brief pulses of L-glutamate or agonist are applied as concentration jumps to outside-out patches excised from extrajunctional membrane of locust (S. gregaria) muscle, openings of channels gated by qGluR appear transiently before the qGluR in the patch desensitize (Dudel et a f . , 1988, 1990; Standley et al., 1993). Dudel et al. (1988) found two kinetically distinct receptors; that is, qGluR which gate L-channels, with long open rimes similar to those reported in the above equilibrium studies, and qGluR which gate s-channels with open times which are brief (< 0.5 ms). A characteristic of many patches used in the concentration jump studies is that qGluR channel activity declined with time, and in some of these patches L-channels appeared to convert to I ) channels before channel openings disappeared. Usherwood (1989) suggested that this change might result from patch deterioration (i.e. rundown) and, therefore, the s-channels might be artefactual. Patches excised from locust muscle do not always contain s-channels, and rundown is not always seen. For example, in a recent study by Standley et a f . (1993) a patch of locust (S. gregaria) muscle membrane containing at least five qGluR was exposed to a continuous 0.33 Hz train of 50 ms pulses of 3.3 x lop3 M L-glutamate for 17 min. The L-channel currents recorded from this patch were similar at the beginning and end of the experiment. Not unexpectedly, the rate of activation of qGluR during a concentration jump is proportional to the concentration of L-glutamate applied to the membrane patch. The channel opening rate estimated from the rising phase of the qGluR cumulative current is c. 5 x 105/ds, which compares with a value of 1 0 4 / ~ / sobtained by fitting the megaohm seal data to Model 1 (Kerry et a f . , 1988a). However, the apparent KD for L-glutamate, derived

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from concentration jump experiments (Standley et al., 1993) and from gigaohm seal equilibrium studies of Bates et a f . (1990) is c. 1.5 x M, that is, about 10 times less than that obtained in the megaohm seal equilibrium studies. 3.2.3 Desensitization of quisqualate-sensitive glutamate receptors When Dudel et al. (1990) applied a slowly rising concentration ramp to outside-out patches of locust (S. gregaria) muscle membrane, the qGluR that they contained were desensitized without channel openings. Therefore, it follows, that desensitization of this receptor can proceed from a closed channel state. In contrast, desensitization of nAChR is usually modelled with the desensitized state(s) proceeding from the open channel state (Dudel et al., 1992). Outside-out patches of locust muscle membrane rarely contain more than 15 qGluR. This observation is commensurate with the low population density of extrajunctional qGluR (Cull-Candy and Usherwood, 1973). The time course of the response of a membrane patch to L-glutamate was best evaluated, therefore, by averaging the currents obtained in response to many (i.e. usually 30-40) concentration jumps or pulses of L-glutamate. These averaged data showed that after reaching an initial peak, within c. 10 ms, the qGluR current declined, presumably as the qGluR desensitized. The averaged current for L-channels declined biphasically , with time constants c. 3 ms and c. 50 ms, although these values varied from patch to patch (Standley et al., 1993). In contrast, the averaged current for s-channels declined monophasically with a time constant of c. 3 ms (Dudel et a f . , 1988, 1990). A major difficulty has been encountered in attempts quantitatively to relate the decay of the averaged currents of L-channels with onset of desensitization, namely that when the concentration of L-glutamate was raised the duration of the averaged current declined. This is opposite to expectation, because the rate of desensitization of a transmitter receptor might be expected to have increased under this condition. The increased duration of the averaged qGluR current is related to the complex kinetics of this receptor. It can be seen from Model 1 that when the concentration of agonist is raised the multiple-liganded open states become more highly populated. Since desensitization proceeds from an unliganded closed state, it follows that the multiple-liganded states will slow the rate of desensitization onset, because the receptor must first return to its unliganded closed state before desensitization is possible (Standley et a f . , 1993). Recovery from desensitization has been investigated using double pulses of agonist (Dudel et al., 1988, 1990; Standley et al., 1993). In the case of qGluR which gate s-channels, recovery is very rapid, that is, complete within a few milliseconds. In the case of qGluR which gate L-channels, recovery takes many seconds and is biphasic. The latter observation suggests


32 1

that desensitization is not a single-step process. This conclusion receives support from data obtained from both megaohm and gigaohm seal recordings, in which qGluR were equilibrated with agonist without pretreatment of membrane patches with concanavalin A; openings of qGluR were rarely seen under this condition. At least two desensitization reactions involving the binding of agonist to qGluR must be invoked to account for the lack of cyclical recovery in these studies. Following the demonstration by Dudel et al. (1990) of biphasic recovery from desensitization for locust muscle qGluR, a qualitatively similar phenomenon was reported for vertebrate hippocampal NMDA and non-NMDA receptors (Colquhoun et al., 1992; Lester and Jahr, 1992). Attempts to model data obtained from concentration jump studies of qGluR have had to contend with: (1) the apparent (about 10-fold) lower sensitivity of these receptors in gigaohm seal recordings (equilibrium and concentration jump) from outside-out patches (Dudel et al., 1988; Bates et al., 1990) compared with megaohm seal recordings from cell-attached patches (Kerry et al., 1988a); and (2) the apparently higher activation rate of qGluR in the concentration jump studies. Model 1 provides a good prediction of the data obtained from megaohm seal studies but it does not fit the concentration jump data. Somewhat surprisingly, the best fit to the latter is obtained with a simple linear model (Model 2) with three closed states. two of which are liganded, and a single open state! kdl/KEL

DA*\DA&I

kdJKES ak,,

2ak,,

c+ CA kOnk

akon

2k,,"/KB

h

\~/K,,OA?

CA~

(2)

where: D is a desensitized state of the receptor-channel; KB = 7 6 9 2 / ~ KO, ; the equilibrium constant for opening of the (biliganded) receptor-channel = 1.8; KES, the L-glutamate binding constant for the short-lived desensitized state = 4.7 x l O P 7 / ~ ; KEL, the L-glutamate binding constant for the longlived desensitized state = 2 1 4 3 5 / ~ ;k,,, the opening rate of the glutamatereceptor channel = 104/dms; kds, the L-glutamate binding rate to the short-lived desensitized state = 1 0 4 / ~ / m s kdl, ; the L-glutamate binding rate to the long-lived desensitized state = 2.14Mms; and h = 0.9/ms. Why are qualitatively different models required to fit the equilibrium and concentration jump data? Attempts to answer this question must bear in mind that the two sets of data are not strictly comparable. Model 1 is derived from a rigorous quantitative assessment of p O ,mo, m, and fo over a wide range of L-glutamate concentrations and, as such, embraces what might be termed the fine details of qGluR channel kinetics. Model 2 is simply the best fit to averaged currents obtained in concentration jump experiments and, as such, it does not directly take into account factors such as the dependence of rno on L-glutamate concentration and correlations between open states and closed states of the qGluR channel.

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One explanation for the low activation rate for qGluR estimated from the megaohm seal experiments is that concanavalin A did not completely block desensitization of this receptor. A partial block would lead to a lower than expected fo, which is one measure of the rate of qGluR activation. The high KD estimated from in the gigaohm seal data is more difficult to explain, but might be related to changes (perhaps involving phosphorylation sites on the intracellular domain of qGluR) which take place in qGluR during manufacture of outside-out patches of locust muscle membrane. Alternatively, the use of enzyme to clean the surface of muscle fibres prior to gigaohm seal formation may affect the properties of qGluR, for example, the inability of concanavalin A to bind in a manner which invokes qGluR desensitization block. The discovery by Mathers and Usherwood (1976, 1978) that concanavalin A blocks desensitization of locust muscle qGluR has only recently begun to have a widespread impact in neuroscience, following reports that this lectin also blocks desensitization of some mammalian ionotropic GluR (Mayer and Westbrook, 1987). In an attempt to understand the mechanism underlying this phenomenon, Evans and Usherwood (1985) tested a variety of lectins, with different sugar specificities and/or valencies, on qGluR of locust ( S . gregaria) leg muscle. Briefly, they found that lectins (e.g. pea and lentil lectins) with the same simple sugar specificities (mannose/glucose) as concanavalin A also block desensitization of qGluR, whereas lectins (soybean and wheatgerm) with other sugar specificities do not. Concanavalin A is a tetramer at pH 7, whereas succinyl-concanavalin A is a dimer. Although the later does not block desensitization, prior application of this lectin prevents desensitization block by subsequent applications of concanavalin A. Evans and Usherwood (1985) also found that when concanavalin A is applied to desensitized qGluR, desensitization is not reversed, and that future applications of this lectin, even in the absence of agonist, do not block desensitization. What explanation is there for these findings? One might envisage the following scheme. Let us assume that qGluR is a tetrameric homo-oligomeric receptor, although the argument put forward below is not dependent upon this being the case. Assume that agonist binds to all four subunits and that desensitization requires a conformational change in these structures. Further assume that, in the absence of agonist, a molecule of tetravalent concanavalin A binds to mannose residues on each of the qGluR subunits, thus cross-linking the four subunits. As a result, the conformational change required for desensitization is no longer possible, that is, desensitization is blocked. Finally, assume that it is not possible for concanavalin A to crosslink the subunits of desensitized qGluR for steric reasons, but the lectin can still bind to single subunits. Since crosslinking of all four qGluR subunits is essential for desensitization block, and because the binding of concanavalin A is only very slowly reversible, it follows that subsequent applications of concanavalin A, even to


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resensitized qGluR, will not block desensitization of this receptor. The lack of desensitization block with divalent succinyl-concanavalin A shows that even when two subunits are crosslinked, this is insufficient to cause desensitization block. Interactions of lectins with insect ionotropic GluR is a ripe area for a study in which qGluR of locust muscle could play a pivotal role.

3.2.4 Ion-selectivity of quisqualate-sensitive glutamate receptor channels The ion-selectivity of extrajunctional qGluR of locust ( S . gregaria) muscle has been studied using megaohm seals (cell-attached patches), with muscles pretreated with concanavalin A, and following equilibration of membrane patches with lop4 M L-glutamate (Kitts and Usherwood, 1988). The alkali metal ions Li, Na, K, Rb and Cs were all highly permeant when these were the major positive charge carriers in the patch pipette saline. The relative conductances of these cations were Li+ < Na+ < Cs+ = K+ < Rb+. The qGluR channel is permeable to NHZ, but impermeable to organic monovalent ions such as tetramethylammonium, guanidinium and choline. High concentrations (e.g. 30 mM) of divalent cations cause channel block. It appears that the qGluR channel combines a high conductance with a restricted ion-selectivity, based on ionic charge and size, the conductance being dependent upon the dehydration energy of the ionic species. Extrajunctional qGluR channels seem to differ from those of junctional GluR of locust muscle in terms of their permeability to divalent cations. Anwyl and Usherwood (1974) showed that GluR of the postsynaptic membrane of locust metathoracic extensor tibiae muscle gate channels permeant to calcium when exposed to saline containing L-glutamate, although the channels are blocked when the sodium in the saline has been entirely replaced by calcium (Kitts and Usherwood, 1988). In contrast, Kits and Usherwood (1988) showed that the calcium permeability of the qGluR channel is low, although, like the postjunctional GluR channel, this channel is blocked by high concentrations of calcium. The apparent differences in calcium permeability of channels gated by qGluR on the one hand and postjunctional GluR on the other hand does not imply, necessarily, that qGluR and quisqualate-sensitive postjunctional GluR are different in this respect. The postjunctional GluR population is not pharmacologically homogeneous; ibotenate-sensitive and aspartate-sensitive GluR (both of which gate cation-selective channels) are present (Fig. l ) , in addition to quisqualate-sensitive GluR (Gration et al., 1979), and one or both of these GluR subpopulations may account for the calcium gating seen during application of L-glutamate.

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3.2.5 Non-competitive receptors

antagonism

of

quisqualate-sensitive

glutamate

Single-channel studies of the interactions of philanthotoxin-433 (PhTX-433) (Clark et al., 1982; Karst and Piek, 1991), phencyclidine (Idriss and Albuquerque, 1985), (+)-tubocurarine (TC) (Kerry et al., 1987), trimetaphan (Ashford et al., 1987), chlorisondamine (Ashford et a/., 1988), argiotoxin636 (ArgTX-636) (Kerry et al., 1988b) and ketamine (Macdonald et al., 1992) with either postjunctional GluR or extrajunctional qGluR have provided quantitative information on non-competitive antagonism of these receptors in S . gregaria and L. migratoria. 3.2.5.1 Channel block by (+)-tuboururine It has been pointed out earlier that po of the qGluR channel increases when the concentration of Lglutamate is raised. However, in the presence of 5 x lop5 M T C this effect of agonist was eliminated (Kerry et al., 1987). Correlations between successive openings and closings of the qGluR channel, which characterize the single-channel activity elicited by L-glutamate, were weakened by TC. This phenomenon is predicted by Model 1 if the blocking agent has a low unblocking rate (see below). However, no attempt has yet been made quantitatively to analyse qGluR channel block by T C using Model 1. Instead, a simple scheme (Model 3) for channel block has been used, viz:

where C is the closed channel of GluR, 0 is its open channel and OB is its open but blocked channel. The channel gating mechanism is represented as a pseudo-unimolecular process with k,, as the (agonist-concentration dependent) opening rate and k, as the closing rate. k,, is the dissociation rate of the blocker-channel complex. The estimated dissociation constant for T C is 1.75 p~ (at V, = -100 mV). The rate of association of T C with the qGluR channel at V, = -100 mV is estimated to be 8.74 x 1Ohls1M,whereas that for dissociation, estimated directly from the single channel data, is 15.31s. Unfortunately, the megaohm seal technique that was employed in these studies did not enable the effect of T C to be studied over a wide voltage range, so it was not possible to determine whether block of the qGluR channel by this compound is voltage-dependent; although on the basis of a limited study of voltage-dependence Kerry et al. (1987) claim that it is not. 3.2.5.2 Antagonism by phencyclidine, ketamine, chlorisondarnine and trimetaphan Phencyclidine is a general anaesthetic and a hallucinogenic agent, which interacts with an open channel state of postjunctional GluR of locust ( L . migratoria) excitatory nerve-muscle junctions (Idriss and


325

Albuquerque, 1985). This seemingly simple channel block contrasts with the action of ketamine on postjunctional GluR and qGluR. Ketamine is also a general anaesthetic, which causes open channel block of nAChR of ) vertebrate skeletal muscle (Maleque et al., 1981), at doses (c. 20 p ~ close to those that cause clinical anaesthesia. Ketamine (50 PM) increased the rate of onset of desensitization of qGluR of locust ( S . gregaria) skeletal muscle. an effect which was abolished by pretreatment of this muscle with concanavalin A (Ashford et al., 1989). Ketamine also reduced the amplitude of the excitatory postsynaptic current (EPSC) recorded from voltageclamped muscle fibres, but the rise time of the EPSC was unaffected and the effect on EPSC amplitude was voltage-independent. The decay of the EPSC is normally monophasic, but during ketamine treatment it became biphasic. This is indicative of open channel block, but the biphasic decays were not influenced by voltage. Ashford et al. (1989) concluded that ketamine interacts mainly with a closed channel state of the postjunctional GluR to prevent channel gating, although it may also cause open channel block. In subsequent single channel studies (using a megaohm seal technique) of ketamine action on extrajunctional qGluR of locust ( S . gregaria) muscle, Macdonald et al. (1992) showed that lop8 M ketamine decreased p o , through open and closed channel block; f o and mo were both reduced. They also showed that this anaesthetic dissociates only slowly from its open channel blocking site on qGluR. In subsequent studies with ketamine, Macdonald el al. (1993) studied the effects of high pressure on qGluR channel block by ketamine. The major objective of this work was to attempt more clearly to define the antagonism, that is, is it steric or allosteric? In the former case one would expect the blocking molecule to bind to the channel, thereby occluding it and reducing its conductance to zero. Allosteric blockers are considered to elicit a conformational change in transmitter receptor proteins and, thus, indirectly reduced the conductances of the channels gated by these receptors. In the studies of Macdonald et al. (1993), pressure was applied with helium gas and recordings were undertaken in a pressure chamber at atmospheric pressure, at 10 MPa pressure and at 50 MPa pressure. In the absence of ketamine, the kinetics of qGluR were unchanged at 10 MPa, but p o and rno were reduced at 50 MPa compared with controls (obtained at atmospheric pressure). At atmospheric pressure, ketamine reduced p o as before (Macdonald et al., 1993), but this effect was reversed by raising the pressure to 10 MPa. This result implies that antagonism of qGluR by ketamine involves a large increase in enthalpy and, therefore, a conformational change in qGluR. In other words, non-competitive antagonism of qGluR by ketamine is probably allosteric. The complex effects of trimetaphan on the EPSC of locust ( S . gregaria) muscle have been interpreted on the basis of enhancement of desensitization of postjunctional GluR and open channel block of these receptors (Ashford et al., 1987). Single channel studies of extrajunctional qGluR made using

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326 0

I

/ /

/

I I

*

GLUTAMATE

@ d

POTENTIATION ANTAQONISM

I

r-

I

OUT

IN


327

patch pipettes containing M L-glutamate with and without the antagonist (at lop4 M) showed that channel openings were interrupted by brief closings when the antagonist was present. This effect of trimetaphan resembles that of many open channel blockers, for example local anaesthetics and their derivatives, on nAChR (Neher and Steinbach, 1978; Ogden et al., 1981). Chlorisondamine action on locust (S. gregariu) muscle EPSC and extrajunctional qGluR suggest that this compound (a diquarternary amine tetrachloroisoquinoline) interacts with closed and open channel states of qGluR (Ashford et al., 1988). Not surprisingly, for a compound carrying a net positive charge, open channel block by chlorisondamine increased with hyperpolarization. However, at membrane potentials of about - 120 m V there was a reduction in the effectiveness of this compound, possibly because the molecule is swept through the channel gated by qGluR (see below).

3.2.5.3 Antagonism by polyamine amides Polyamine-containing toxins isolated from venoms of certain spiders (e.g. araneids) and the venom of a wasp, Philanthus triangulum, are powerful non-competitive antagonists of GluR (Fig. 2). These aryl amines or polyamine amides are low molecular weight compounds (often < 1 kDa) containing one or more amino acids, and are polycations at physiological pH. They contain an aromatic chromophore and a polyamine backbone, and most terminate in a primary amino or guanidino group (Fig. 2). A large number of natural polyamine amides have been described and over 100 analogues have been synthesized. Studies of the sites and modes of action of these compounds with respect to their antagonism of qGluR have mainly concentrated on the natural FIG. 2 Top: the generalized structure of a polyamine amide toxin (see text for further details). When polyamine amide toxins of spiders (e.g. argiotoxin) and parasitic wasps (philanthotoxin) enter a prey insect they target its glutamatergic nerve-muscle junctions. Site I: the toxins potentiate and non-competitively antagonize the postjunctional GluR and the extrajunctional muscle qGluR. Site 2: the presence in the venoms of high concentrations of L-glutamate and the occurrence (presynaptically on motor axon terminals) of GluR which enhance spontaneous and evoked release transmitter L-glutamate leads to an elevation of 1.-glutamate concentration in the synaptic cleft which assists the action of the toxins. Site 3: the toxins may block the high affinity uptake of L-glutamate into axon terminal and glia. which will contribute to the elevation of the concentration of this amino acid in the synaptic cleft. (From Usherwood (1991b). Bottom: summary of interactions of PhTX-343 with postjunctional GluR and extrajunctional qGluR of locust skeletal muscle. The toxin binds to four sites on these receptors; two are on the extracellular domains of the receptors, one is in the receptor channel, and the fourth site is on the intracellular domains of the receptors close to the internal orifice of the channel. Three of the sites (0)are involved in non-competitive antagonism of these receptors: binding of toxin the other site (0)allosterically potentiates the receptors.

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products, argiotoxin-636 (ArgTX2-636), where 636 is the molecular weight of the toxin, and philanthotoxin-433 (PhTX-433) (or its almost equipotent, synthetic analogue philanthotoxin-343 (PhTX-343)), where the numerals refer to the number of methylene groups connecting the amine moieties of the polyamine backbone. Discoveries of polyamine-containing spider and wasp toxins, apparently designed by nature specifically to antagonize transmission at glutamatergic nerve-muscle junctions of prey insects, have resulted in a flurry of publications on insect muscle glutamate receptors. There is a large number of up-to-date reviews of work in this area to which the reader is referred (Usherwood, 1987a, b; 1988, 1989, 1991a, b; Jackson and Usherwood, 1988; Adams, 1988; Pick et al., 1988; Nakajima et al., 1988; Usherwood and Blagbrough, 1989a, b, 1991; Blagbrough and Usherwood, 1992; Chiles et al., 1992; Nakanishi et al., 1992; Scott et al., 1993). Here I will summarize only the salient properties of these toxins. Polyamine amides interact with insect qGluR in at least three ways: (1) to induce potentiation, by binding to a site on the extracellular domain of this receptor (Usherwood, 1988; Kerry et al., 1988b; Karst and Piek, 1991); (2) to antagonize qGluR, by binding to a closed channel state of this receptor (Magazanik et al., 1986; Kerry et al., 1988b), presumably also at an extracellular site; and (3) to antagonize qGluR through open channel block (Usherwood et al., 1984; Bateman et al., 1985; Magazanik et al., 1986; Kerry et al., 1988b). The relative apparent affinities of these three proposed binding sites for locust qGluR (Fig. 2) are; potentiation site > closed channel block site > open channel block site (Kerry et al., 1993), although closed and open channel block of blowfly lava1 muscle glutamate receptors by ArgTX-636 have similar K,s (Magazanik et al., 1986). There may be a fourth binding site for these toxins on qGluR. When PhTX-343 is injected into locust ( S . gregaria) muscle fibres it antagonizes qGluR apparently by binding to an internal site on this receptor, possibly close to the internal orifice of the channel gated by this receptor (Brundell et al., 1991) (Fig. 2 ) . It remains to be determined whether extracellularly applied toxin can gain access to this internal site by passing through the channel gated by qGluR (Usherwood and Blagbrough, 1989a). Despite the progress that has been made in recent years in understanding potentiation and antagonism of qGluR by polyamine amides it is clear that further studies are required if the binding sites for these toxins are to be identified unequivocally. The recent synthesis of potent photosensitive, radiolabelled analogues of PhTX-343 (Goodnow et al., 1991) and ArgTX636 (Nakanishi, personal communication) could have a major impact in this respect, but progress will be more rapid when the subunits of qGluR have been cloned and structurally characterized. Structure-activity studies of polyamine amides with respect to their interactions with insect muscle GluR have broadly involved either analogues

329


of PhTX-343 (Bruce et a f . , 1990) and PhTX-433 (Karst et al., 1990; Karst and Piek, 1991; Benson et al., 1992), natural analogues of ArgTX-636 (e.g. Budd et al., 1988; Grishin et al., 1989) or hybrids of ArgTX-636 and PhTX433 (Blagbrough and Usherwood, 1992). A number of general conclusions about the structural factors which influence potency have emanated from these studies, for example, potency is influenced by the number of positive charges carried by the polyamine moiety; potency is increased when the toxin has a terminal guanidino group; hydrophobicity of the aromatic moiety is an important factor in potency determination, and potency is increased by the presence of a hydrophobic side chain (or side chains) as in some synthetic PhTX-343 analogues.

3.3

DEVELOPMENTAL STUDIES OF SKELETAL GLUTAMATE RECEPTORS

A comparison of glutamate receptor channels of larval and imaginal muscles of the mealworm (T. molitor) has raised some interesting questions about possible changes in the properties of these macromolecules during development of this insect. According to Saito and Kawai (1987), channels gated by L-glutamate in larval muscle have a conductance of c. 15 pS whereas those of imaginal muscle are about 30 ps. Unfortunately, the markedly different salines (larval muscle saline contained 70 m M NaCI, 30 mM KC1, 2 mM CaC12, 15.5 mM MgCI2, 475 mM sucrose, although in some experiments the MgCI2 was omitted; imaginal muscle saline contained 14 mM NaCI, 4 mM KCI, 2 mM CaC12).used in these studies makes it difficult to evaluate the significance of the measured differences in conductance. Also, since p o for the channels elicited from larval muscle showed only a weak dependence on L-glutamate concentration, one might reasonably question their identification with GluR. In a parallel study (Saito and Kawai, 1985), these authors found H-receptors and D-receptors for L-glutamate on T. molitor imaginal muscle fibres (only the D-receptors were found on larval fibres) (see below).

3.4 IBOTENATE-SENSITIVE GLUTAMATE RECEPTORS

OF SKELETAL MUSCLE

EXTRAJUNCTlONAL MEMBRANE

The significance of the discovery by Lea and Usherwood (1973) of Hreceptors or iGluR on locust ( S . gregaria) leg muscle which gate chlorideselective ion channels is still not clear. Outside-out patches of membrane excised from metathoracic extensor tibiae muscle fibres of the locust ( S . gregaria) have been studied using the liquid filament switch technique of Franke et al. (1987) for rapid application and removal of (within 1 ms in both cases) drugs and ions (Dudel et a [ . ,

330

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1989). When the liquid filament contains L-glutamate, cation-selective (100-150 pS) and anion-selective (25 pS) ion channels are gated in a patch. The latter arise through activation of iGluR. A single outside-out patch contains a large number (> 140) of iGluR. The iGluR channels are not gated by L-quisqualate, NMDA and kainate, but they were gated by DLibotenate. Interestingly, a high concentration (lo-* M) of L-aspartate also gates the iGluR channel. Unlike the GABA receptor channel of locust muscle, that of the iGluR is slightly permeable to methylsulphate. Lea and Usherwood (1973) had previously arrived at this conclusion following their studies of extrajunctional H-receptors of locust (S. gregaria) leg muscle using ionophoretic application of drugs and intracellular recording. The reaction kinetics of the iGluR channels seemed to be slower than their qGluR counterparts. Whereas maximum activation of the latter occurred within 1 ms of agonist application to a patch, the maximum current resulting from iGluR activation was reached only after 2&50 ms. Desensitization of iGluR was slow in its onset and incomplete. This contrasts with the rapid and prolonged desensitization of H-receptors seen in ionophoretic studies of locust muscle (Lea and Usherwood, 1973). Of additional interest was the observation that the current generated by iGluR channels decayed only slowly (time constant c. 50 ms) after termination of an agonist pulse (Dude1 et al., 1989).

4 Glutamate receptors of visceral muscle

The GluR of the superior longitudinal muscles of locust ( L . migratoria) hindgut appear to be remarkably similar to those of skeletal muscle (Dunbar and Piek, 1983); a similarity which extends to the presence of D- and Hreceptors on extrajunctional membrane of these muscles. Desensitization of junctional GluR is blocked by concanavalin A and responses to L-glutamate are antagonized by PhTX-433. Channel openings of 135 pS conductance have been recorded from patches of cockroach hindgut muscle fibres using a megaohm seal technique with patch pipettes containing L-glutamate (Kits el al., 1984). The muscle fibres were pretreated with concanavalin A to inhibit GluR desensitization. Unfortunately, technical difficulties precluded the adoption of the rigorous recording and analytical procedures devised by Kerry et al. (1986) for their single-channel studies of locust skeletal muscle qGluR, so quantitative comparisons of data obtained from the two systems are not possible. However, qualitatively, there are sufficient similarities between the single channel properties of GluR of insect visceral and qGluR of skeletal muscle to suggest that the same protein is expressed in both tissues.


5

33 1

Metabotropic glutamate receptors

The possible involvement of cyclic nucleotides as intracellular messengers for some actions of L-glutamate on insect excitable tissues has been raised on a few occasions. Robinson et al. (1982) demonstrated adenylate and guanylate cyclase activation by L-glutamate for membrane preparations of larval dipteran (Sarcophaga sp. ; Lucilia sericata) body-wall muscles. Adenylate cyclase activity was stimulated either in the presence of, or absence of, calcium, although a lower concentration of L-glutamate was required in the former case M compared with M). The structurally related amino acids, L-aspartate and L-cysteine sulphonic acid also activated the two cyclases, but at higher concentrations than Lglutamate. Since the concentrations of L-glutamate required to activate both adenylate cyclase and guanylate cyclase were very low, it was proposed that these intracellular messengers might also be activated by spontaneous release of transmitter from glutamatergic motor nerve terminals (Robinson et af., 1982). However, this proposal still remains untested. This work followed a study by Robinson (1980, 1982) on the depolarization and membrane resistance changes of dipteran body wall muscle caused by Lglutamate. In addition to the expected depolarization and conductance increases, presumably resulting from activation of qCluR, there was a second, lower threshold effect of glutamate in which a slow membrane depolarization was accompanied by a conductance decrease, these effects of L-glutamate being mimicked by CAMP. The lower threshold effect of Lglutamate is tentatively analogous with the response to L-glutamate of metabotropic GluR of mammalian CNS (Nicoletti et af., 1986). For this reason the insect GluR responsible will be termed an insect metabotropic GluR (mGluR, Fig. 1). Robinson (1982) also showed that CAMP mimics the effect of L-glutamate in respect of the slow depolarizations and conductance decreases. The pharmacology of mGluR is different from that of qGluR; of particular note is the high potency of L-asparate, an amino acid which is almost inactive on qGluR. In view of this, it might be a particularly useful to employ L-aspartate rather than L-glutamate in future studies of insect mGluR. Since the discoveries of octopamine and proctolin, there have been many suggestions that these compounds might act on GluR of insect CNS and muscle (skeletal and visceral). Yamamoto and Ishikawa (1991) found that 5 PM octopamine potentiated the excitatory postsynaptic potential of larval mealworm (T. rnolitor) ventral longitudinal muscle fibres, possibly through a presynaptic mechanism, but this compound had no effect on the potential generated by glutamate ionophoresis at neuromuscular junctions. However, it reduced the depolarization induced by bath-applied L-glutamate. Although they had assumed from studies of Saito and Kawai (1985) that extrajunc-

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tional membrane of mealworm larval muscle fibres does not contain ionotropic GluR, they concluded that octopamine reduces the sensitivity of the extrajunctional membrane to L-glutamate. An investigation of the pharmacology of this phenomenon indicated that octopamine-induced attenuation of the extrajunctional glutamate response is mediated by cyclic nucleotides. The action of octopamine was mimicked by forskolin, CPTcyclic AMP and 8-bromo-cyclic GMP, but not by a protein kinase C activator, 1,2-oleoylacetylglycerol.The ability of octopamine to stimulate the production of CAMP but not cGMP in various insect excitable tissues is well-documented, with phosphorylation of transmitter receptor proteins being implicated in some cases.

6 Cloning, expression and functional characterization of glutamate receptors

6.1

EXOGENOUS EXPRESSION OF NATIVE RNAS

The Xenopus laevis oocyte technique has been used recently for expression of insect GluR. Fraser et al. (1990) extracted mRNA from leg muscles of the locust (S. gregaria) and injected this into Xenopus oocytes. Receptors for Lglutamate, both qGluR and iGluR, were expressed, but in only a small minority of oocytes. Possibly, the fraction of mRNA encoding GluR that is extracted from locust muscle is very small. Attempts to increase the amount of specific RNA by increasing the quantity of mRNA injected, that is, by injecting RNA of denervated muscles which are known to express a higher than normal population of extrajunctional qGluR (Usherwood, 1969), or by using various stages in the locust life history (embryo, hopper and adult) all failed to increase the number of oocytes expressing GluR. L-Quisqualate evoked an inward current at -60 mV which reversed at about -9 mV. This response was occasionally accompanied by an oscillatory component, not unlike that associated with metabotropic GluR of vertebrate central neurones and may reflect the presence of insect mGluR. Sometimes the quisqualate-induced current could not be fully reversed, an observation which supports the presence of a metabotropic component. However, Lglutamate elicited biphasic currents, but did not induce an oscillatory response. Rapid desensitization often accompanied application of Lglutamate, but this could be inhibited by pretreatment of the oocyte with concanavalin A. The response to ibotenate comprised a smooth inward current with a reversal potential of about -21 mV, that is, the chloride equilibrium potential for an oocyte. L-Aspartate and L-kainate were inactive, even at 1 mM concentration.


6.2

CLONING

AND

STRUCTURAL

333 CHARACTERIZATION

OF

CENTRAL

AND

PERIPHERAL GLUTAMATE RECEPTORS

The isolation and functional characterization of complementary genes encoding a D. melanogaster kainate-selective GluR (DGluR-I) (Schuster et al., 1991, 1993), a GluR expressed in muscle tissue (DGluR-11) (Ultsch et al., 1992) and a putative NMDA receptor (DNMDAR-I) expressed in D . melanogaster brain (Ultsch et al., 1993) have been reported. DGluR-I is a 964-residue protein with a molecular weight of c. 109 kDa. Its transcripts are differentially expressed during D.melanogaster development and, in late embryogenesis they accumulate in the CNS. Hydropathy analysis of the deduced mature DGluR-I protein revealed four putative membranespanning segments, an arrangement which is identical to that of DGluR-I1 subunits. This suggests that both subunits are like ionotropic GluR of mammalian CNS (Sommer and Seeburg, 1992). DGluR-I and DGluR-I1 also exhibit significant sequence homologies with ionotropic GluR subunits of mammalian CNS. The highest overall amino-acid identity (between 41 and 44%) for DGluR-I is seen to the GluRl to GluR4 class of rat ionotropic GluR subunits, whereas DGluR-I1 exhibits only 26% sequence identity to rat ionotropic GluR subunits. Interestingly, the Drosophila DGluR polypeptides also have a small homology (20%) with the rat brain NMDARl (NR1) receptor subunit (Moriyoshi et al., 1991). When DGluR-I cRNA was injected into Xenopus oocytes, homo-oligomeric GluR activated by kainate were expressed. These ionotropic GluR gate channels which are blocked by CNQX and PhTX-433. They are insensitive to AMPA, quisqualate, NMDA, ibotenate and L-homocysteate at 1 mM or greater concentration. Somewhat perplexingly, they are not activated by Lglutamate; responses to very high concentrations (> 10 mM) of this amino acid were seen in oocytes injected with DGluR-I cRNA, but these currents were considered to be non-specific since they were not blocked by CNQX and they also occurred in oocytes, which had not been injected with the cRNA. When DGluR-I1 was expressed in oocytes responses were elicited by L-glutamate and L-aspartate, but only when very high concentrations (10 mM) of these amino acids were applied. The low sensitivity to these amino acids possibly reflects the absence of complementary subunits. Three partial clones (Locl-Loc3) of putative GluR subunits of the locust (S. gregaria) have also been described (Usherwood et al., 1993). These exhibit significant sequence identity with the two D. melanogaster clones. Assuming a comparable length to that of rat GluRl subunit, Locl is deduced to be missing c. 100 amino acids of the C-terminus and c. 200 amino acids at the N-terminus.

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7 Summary

Considerable progress has been made over the past decade or so in gaining an understanding of the biophysical and pharmacological properties of insect GluR, in particular qGluR, and studies of locust muscle continue to provide basic information on transmitter receptor function. The future of research in this area is very exciting, but depends greatly on the cloning and structural characterization of more insect GluR subunits. Although information on the subunit compositions of ionotropic receptors of vertebrate CNS and muscle might lead one to anticipate that insect GluR are hetero-oligomers, available evidence, albeit of a rather scant nature, does not exclude the possibility that at least some are homo-oligomers. Information on the genes for these subunits will raise opportunities for transgenic studies, the equivalent of which will be difficult, perhaps impossible in mammals, and also for the exploitation of Drosophila genetics further to manipulate GluR. Cloning of insect GluR subunits will also make possible much greater insight into the distributions of GluR in insects and their developmental relationships. The recent discovery of presynaptic NMDA receptors at locust nerve-muscle junctions (L. Magazanik, personal communication) raises some interesting issues in comparative neuroscience, and also affords further opportunities to employ insect preparations as models for basic neuroscience research. The question of how the insecticide industry might best exploit our knowledge of insect GluR remains unanswered. The insect excitatory nerve-muscle junction seems to offer an enticing, unexploited target site, yet despite the wealth of seemingly relevant information on this system it still remains untouched.

Acknowledgements

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Index Tables in bold; Figures in italic. Abdominal ganglia, Arthropoda, 17, 18, AMPA (Amino-3-hydroxyl-5-methyl-419 isooxazole propionic acid), 312. 313,333 Accessory glands, juvenile hormone, Amphibians, 131, 168-9, 197, 253 218,219,246, 247,248 Anaesthetics, 324, 325 Accessory planta retractor motor Anaphylaxis, 122 neurons (APR), 242 Androctonnus australis, 175 Acetylcholine systems, 310 Annelida, 58 Acetylcholinesterase, 72, 244 Antennae, 29. 33, 43, 44.46, 234 Acheta, 29, 39 Antenno-glomerular tract, Arthropods, domesticus, 129-31, 139, 140, 141, 46 147, 150, 159, 169, 224 Antheraea polyphemus, 230 Acridid, 25 Antherix variagata, 135 Acridoidea, 33 Antibacterial proteins, 162 Acroneuria, 141 Antidiuretic hormones, 169, 172. 173 Activation, Bacillus thuringiensis, 287-8 Antp gene, Arthropoda, 78-9 Acyrthosiphon pisum, 140, 141 Anurogryllus muticus, 156 Adenosine 3'5'-cyclic monophosphate Apis mellifera, 45, 5&1, 55, 56, 314 eicosanoids, 117, 131, 171, 173, 174 Aplysia, 179 glutamate receptors, 331, 332 Apomorphy, Arthropoda, 5 juvenile hormone, 223 Apterygote. 81 Adenosine triphosphate, gut, 284, 292 Apulmonata, 73 Adenyl cyclase, 131, 173, 177, 183, 223, Aquatic invertebrates, eicosanoids, 135 Arachidonic acid and other PUFAs, 116, 331 117,118, 119-29, 120,121,123-7 Adephagan beetles, 50 insects, 128-9 Adipokinetic hormones, 177-8, 177, 185 advances in biochemistry, 187-8, Aedes aegypti, 134, 169,170, 172 196 Aeshna, 26 biosynthesis, 136-47, 138, 141, 142. Agelenids, 72 143 Alkyl-substituted glu tamates, 316 immunity, 165, 166-8, I66 A lpheus heterochelis, 65 neurophysiology, 179 Aminergic neurotransmitters, 12 occurrence in insect lipids, 131-6, Amino-3-hvdrox~l-5-methvl-4-isooxazole 132, 134 propionicacid (AMPA), 312, oxygenation, 129-31 313,333 peptide hormone, 223 Aminoacetophenone, 183 reproduction, 149-51, 152-3, 160, Ammonium, 323 161 AMP see Adenosine 3'5'-cyclic secretion rate, 171, 172, 173, 174 monophosphate . . thermobiology, 176

344

Arachidonic acid and other PUFAs (cont.) mammals, 119,121, 122-7,123-7 Arachnida, 12, 14, 52, 72-3, 80 suboesophageal ganglion, 73-4 supraoesophageal ganglion, 74-6, 75, 76 Archimantis, 41 Areneae, 72, 73, 76,326, 327 Arenophilus, 58 Arginine vasotocin, 161 Argiotoxin-636, 324,326, 327-8, 329 Aromatic compounds, juvenile hormones, 254 Artemia salina, 83 Arthropoda, 161 see also Chelicerata, Homology, Mandibulata Arylphorin, 235-7,236, 238, 239,244 Ascending neurones, 36, 37, 41, 44,45, 55 Aspartate, 285, 331, 332, 333 central nervous system, 311, 312 skeletal muscle, 323, 324, 330 Aspirin, 150, 157, 176, 183 ATPase (Adenosine), gut, 284, 292, 294, 295 Attacins, 162 Auditory organs, Arthropoda, 30-1, 33, 36

Bacillus thuringiensis, endotoxin, 275-7, 298-9 classification of, 277-8, 278 insect gut, 282 Coleoptera, 284-5 Diptera, 284 Lepidoptera, 282-4,283 mechanism of action, 2856,286 activation, 287-8 cell lysis, 2 9 1 4 peritrophic membrane, 288 pore formation, 291 receptors, 288-90 solubility, 2 8 6 7 models for the mechanism of pore formation, 294-6,295,297 ‘penknife model’, 2968,297 umbrella model, 297, 298 structure Cry toxins, 279-80,280,281 Cyt toxins, 280-2 use of, 278-9

INDEX

Bacterial infection, 162-3 Baculovirus, 246, 249 Balanus balanoides, 161 Barium, 293 Barnacle, 161 Barytettix psolus (Mexican grasshopper) 35 Bathus occitanus, 175 Bauplan, Arthropoda, 5, 12, 13, 79, 80 Chelicerata, 71, 74 Crustacea, 68 Insecta, 18, 24, 54, 55 Beetles, 40, 45 Behavioural fevers, 174-5 Binding proteins, juvenile hormones, 2467 Bioassay, 181 Biochemistry, eicosanoids, 186-7 control experiments, 19&4,191-3 molecular biology, 197 phospholipase A2 activity, 187-8 tobacco hornworm tissues, 188-90, 189 control experiments, 19&4, 191-3, 195 fat body preparation, 194-7 Biological control see Bacillus th uringiensis Biosynthesis, PUFAs, 136, 1 8 3 4 biosynthesis of C20 PUFAs, 140, 142, 143, 144-7 de novo biosynthesis, 13940, 141, 142 Lepidopterans, 136-7, 138 mosquitos, 137 Biotechnology, 279 Biphasic response, glutamate receptors, 312,325, 332 Bipolar neurones, 59 Bivalves, 161, 169 Bladder, 197 Blastokinesis, 224 Blattaria, 26 Blattella germanica (German cockroach), 140, 160, 238 Blattodea, 82 Blood-clotting, eicosanoids, 122, 184 Blood-feeding, ticks, eicosanoids, 181-2 Blood flukes, 180-1, 181 Blowfly, 49, 131 Bombyx, 26 mori eicosanoids, 156, 157-8, 159, 159, 177-8

INDEX

juvenile hormones, 216, 2367,236, 238 Boophilus microplus, 181-2 Brachyura, 66 Bradykinin, 182 Bradynotes obesa, 141 Brain, Arthropoda, 2, 6, 7, 80-1 Chelicerata, 72 Insecta, 55-6 Myriapoda, 59 Brassica oleracea, 146 BR-C protein, 252 Bridge, Arthropoda, 74, 75 Bromophenacyl ester, 149 Brown rice planthopper, 184 Bt see Bacillus thuringiensis Bufo marinus, 169 Bumblebees, 175 Buprofezin, 184 Bushcrickets. 33 C-terminal Bacillus thuringiensis, 279, 280, 286, 287,288 juvenile hormone, 247 steroid hormone, 219-20,220 C18 PUFAs, 119,120, 129, 132-3 see also Linoleate, Oleic acid C20 PUFAs, 116, 119,120, 129 occurrence in lipids, 131-6, 132, 134 see also Arachidonic acid, Eicosapentaenoate, Homoylinolenic acid Caelifera, 30 Calcium Bacillus thuringiensis, 293 eicosanoids, 173, 174, 186, 187-8 glutamate receptors, 323, 331 peptide hormone, 223 Calliphora erythrocephala, 49, 55, 56, 131 vicina, 52 Calmodulin, 223 Calpodes ethlius, 236 Cambarus bartoni, 174 Cancer, 66 Cannula punctata, 141 Carausius morosus, 27 Carcinus, 67 menas, 51 Catecholamine, 65, 219, 222 Caterpillars, 215

345

Cecropins, 162 Cell-cell interactions, Arthropods, 37 Cell death, Arthropoda, 18, 19 Cell expression, Arthropoda, 31 Cell lysis, Bacillus thuringiensis, 2 9 1 4 Cell migration, Arthropoda, 31 Cellular immunity, 162, 163-8, 198, 277 Centipedes, 59 Central body, brain, Arthropoda, 2, 3 Chelicerata, 71, 74, 75 Crustacea, 69 Insecta, 33, 46 Central nervous system, 3 1 W , 311 see also Skeletal system Cercal receptors, Arthropoda, 39 Cercarial penetration, 180 Cercus, 29 Cerebral ischaemia, 310 Cesium, glutamate receptors, 323 Channel gating kinetics, extrajunctional glutamate receptors, 316 agonist concentration jump studies, 319-20 desensitization, quisqualate receptors, 320-3 ion-selectivity, quisqualate receptors, 323 non-competitive antagonism of quisqualate-sensitive glutamate receptors, 324 antagonism by phencyclidine, ketamine, chlorisondamine, trimetaphan, 3247,326 antagonism by polyamine amides, 327-9 channel block by (+)-tubocurarine, 324 quisqualate-sensitive glutamate receptors, 3 1 6 7 single channel studies, 317-9 structure-activity studies, 3 1 5 4 Chelae, Crustacea, 65, 66 Chelicerata, 1, 2, 3, 69-71 arachnida, 70-1, 72-3 compared to Insecta, 44 phylogeny, 80,82 ‘primitive’ chelicerata, 71-2 suboesophageal ganglion, 73-4 supraoesophageal ganglion, 70,74-6, 75, 76 visual systems, 77 Chemical mating factor, 147, 149

346

Chemokinesis, 122 Chemoreceptors, Arthropoda, 67 Chemotaxis, 122 Chilopoda, 26, 57, 58, 62, 77 Chironomus tentans, 219, 221, 243 thummi, 238 Chitin, gut, 226, 283, 288 Chitinase, 288 Chloride, 168, 312 Chlorisondamine, glutamate receptors, 324,327 Choline, 168, 310,323 Chordotonal organs, Arthropoda, 28, 30, 31, 62, 66 Chorthippus biguttulus, 36 Chortoicetes terminifera. 20 Chromatin, 221,228, 229, 252, Chromatography, 151, 158, 181 see aEso Gas-chromatographic massspectrometric analysis, Gas-liquid chromatography, Highperformance liquid chromatography Chrysopa carnea, 142 Circadian rhythm, 147 Cladistic analysis, 8, 13 Classification, Bacillus thuringiensis, 277-8,278 Claws, Crustacea, 64 Cloning, glutamate receptors, 332, 333, 334 Co-factors, eicosanoids, 194-7 Cobalt, 20 Cockroach see Periplaneta americana Coelomic cavities, Arthropoda, 44 Coleoptera Bacillus thuringiensis, 276, 277, 278, 282,2865,286 eicosanoids, 128, 142, 183 homologous structures, 26, 47, 54, 82 Colicin A, 298 Colorado potato beetle, 237, 238,285 Columnar cells, gut, 282, 283,283, 285, 286, 2889, 292, 2934,295 Commissures, Arthropoda, 14 Complement proteins, eicosanoids, 162 Compound eye, Arthropoda, 77 Concanavalin A, 313,322 skeletal muscle, 316, 317, 319, 321, 322-3 visceral muscle, 330

INDEX

Connectives, Arthropoda, 14 Continuity, criterion of, Arthropoda, 13-14 Convergent evolution, 11, 69 Coptotermes formosanus, 141 Corpora allata, juvenile hormone, 213, 224,241,244 premetamorphic, 215,216, 217 Corpora pedunculata, Arthropoda, 71, 74 Costelytra zealandica, 285 Coupled GC-mass spectrometry, 117 Courtship behaviour, juvenile hormone, 219 Crab, 67 Cratypedes neglectus, 141 Crayfish, 62, 64, 67, 68, 69, 70 Cricket eicosanoids, 131, 146, 178-9 homology, 25,26, 39, 40 see also Acheta Crista acustica. Arthropoda, 33 Crochets, juvenile hormone, 215,222, 239,240 Crustacea, nervous system, 2, 12, 62, 81, 82-3,82 compared to Insecta, 68-9, 70 development and immunohistochemistry, 8, 9 interneurons, 44, 47, 59, 67-8 motoneurons, 26, 62-5, 63 receptors, 65-7 visual system, 77 Crustacean cardioactive peptide (CCAP), 40, 51-4,53,54,59, 61, 134 Cry proteins, Bacillus thuringiensis, 277, 278,279-80,280,281, 282, 284, 285,298 activation, 287-8 cell lysis, 292, 293, 294 pore formation, 291, 294, 295-6,297, 298 receptors, 28&9 resistance, 290 solubility, 286, 287 Crysomelid beetles, 285, 287 Crysopa carnea, 140 Culex pipiens, 131-2, 132, 133, 134, 137 tarsalis, 132-3, 137 Cupiennius salei, 72, 73, 74-6, 75

INDEX

347

Current concepts of hormone action, Bacillus thuringiensis, 277 219-23,220 homology, 3, 8, 12 steroid hormones, 219-21,220 Cuticle Depolarization, glutamate receptors, homology, 81 312,313 juvenile hormone, 224,22633,227, Dermaptera, 141 233 Dermestus maculatus, 142 Cuticular pegs, Crustacea, 66 A12 Desaturase, 140, 142, 146 Cyano-7-nitroquinoxaline 2,3,-dione Descending contralateral movement (CNQX), 312, 313, 333 detector, Arthropoda, 35 Cyanoprotein, juvenile hormone, 239 Descending neurones, 37,42,46,47.55. Cyclic-AMP-response-element binding 61 protein (CREB), 223 Desert cicadas, 175-6 Cyclic nucleotides, glutamate receptors, Detoxification, 163 331-2 Deutocerebrum. Arthropoda. 43, 44, 45, Cycloheximide, 232 46,57 Cyclooxygenase pathway, eicosanoids, Developmental biology, 3. 4-5, 7-10 119,121, 122,123, 124, 185, 186 fluid secretion rate, 168, 169, 170,170, Developmental genetics, 7, 11 Dexamethasone, 163-7, 164-7 171 Diabrotica undecempunctata howardi, immunity, 162, 166 285 inhibitors, 180, 183 Diacylglycerol (DAG), 173, 177, 188, molecular biology, 197 223 reproduction, 157 Diceroprocta, 176 thermobiology, 176 Dihomo-y-linolenic acid, 119 tobacco hornworm tissues, 194, 19.5, 2,s-Dihydrophenylacetic acid-lactone. 195, 196 183 Cysteate, 312, 314 5,7-Dihydroxy-2-nonyIchrome, 183 Cysteine, 285, 286, 287, 331 2,6-Dihydroxyacetophenone,183 Cyt proteins, Bacillus thuringiensis, .2,4-Dihydroxyacetophenone, 183 277-8, 278, 280-2, 284, 288 1-(2,6-Dihydroxyphenyl)dodecan-I-one. activation, 288 183 cell lysis, 293 Diphtheria toxin, 280 pore formation, 291, 294 Diplopoda, 57, 58, 83 receptors, 289, 290 Diptera Cytochrome P450, 122, I70 Bacillus thuringiensis, 276, 277, 278, Cytoplasmic proteins, juvenile hormone, 282, 284, 285, 286 229-30 eicosanoids, 128, 133, 142, 145-6, 160. Cytosolic juvenile hormone receptors, 187 247-8 glutamate receptors, 331 homologous structures, 8. 14, 17, 26, 47, 54,82 D-channels, 319, 329 juvenile hormone, 214, 216, 229, 233. ‘Dadd’, 128 235 De novo synthesis, 136, 13940, 141, see also Drosophila, Manduca, Musca 142, 144 Diptericins, 162 Decapoda, 62, 6 4 , 6 5 4 , 67, 68, 77 Diuretic hormones, 169, 172, 173 Defence agent, eicosanoids, 182-3 Division rate, Arthropoda, 18 Defensins, 162 DLMs see Dorso-longitudinal muscles Deilephia elpenor, 133, 134 DNA see Deoxyribonucleic acid Dendrites, Arthropoda, 7, 14 Docosahexaenoate, 118, 132, 146 Deoxyribonucleic acid Dodecadienoate, 249

348

Domoate, 312 Donnan effect, 293 Dopa decarboxylase, 227,231,232-3, 233 Dopamine, juvenile hormone, 231,232 Dorsal closure, eggs, juvenile hormone, 215,224 Dorsal glands, eicosanoids, 135 Dorsal unpaired median neurone, DUM cells, Arthropoda, 18, 21, 22, 2 3 4 , 2 6 , 37, 68 Dorso-longitudinal muscles, Arthropoda, 82 Crustacea, 63 Insecta, 2&1,21,22, 25, 26, 37, 40, 47,68 Myriapoda, 5&9,60 Dragonflies, 19, 55 Drosophila homology, 2, 17,79, 81, 83 interneurons, 36, 42, 43, 45, 47, 51 motoneurons, 25, 26 sensory neurons, 28,29, 31,32 hydei, 245 juvenile hormone, 243, 244, 254, 255 epidermis, 229,233,234,235 mechanism, 246,247,253 melanogaster, 11, 50, 52,54, 55, 56, 83 eicosanoids, 128, 142, 145, 146 glutamate receptors, 312,313, 333, 334 juvenile hormone, 216-7,217, 221-2,245 DUM-cells (Dorsal unpaired median neurone), 18,21, 22, 23-4, 26, 37, 68 DUMETi, 24 Dye-iontophoresis, 20 Dysdercus, 26 E74, 252 E75, 252 EBDA (10R. 11s-epoxybishomofarnesyl diazoacetate), 248, 249 Ecdysis essential fatty acids, 128 juvenile hormone, 215-7,217 Ecdysone, 216-8,217,219,220,221-2 see also Juvenile hormone Echinodermata, 161 Ecological significance, eicosanoids, 179-80

INDEX

blood feeding, ticks, 181-2 blood flukes, 180-1,181 inhibitors of eicosanoid biosynthesis, 183-4 predator avoidance, 182-3 Ecydysteroid receptor, 221-2, 251, 253 see also Juvenile hormone EFDA (10,ll-epoxyfarnesyl diazoacetate), 247 Egg case, juvenile hormone, 218 Egg laying, eicosanoids, 130-1, 147-60, 148, 161, 179 EHDA (10R. 11s-epoxyhomofarnesyl diazoacetate), 246, 248, 249 Eicosanoids, 116, 117 comparative physiology, 197-8 desiderata advances in biochemistry, 186-8 mechanism of action, 1854,186 molecular biology, 197 new eicosanoid action, 184-5 fluid secretion rates, 168-73,170, 171, 174 historical perspective, 117-9, 219 immunity, 162-8, 164-7 modulation of lipid mobilization, 177-8, 177, 183 neurophysiology, 178-9 reproduction, 14741,148, 153, 154, 155, 159 thermobiology, 174-6 see also Arachidonic acid, Ecological significance, Manduca sexta Eicosapentaenoate, 118, 119,120, 121, 122,123,124,125, 136 biosynthesis, 146 immunity, 167 lipids, 132, 133, 134 thermobiology, 176 Eicosatetraynoic acid (ETYA), 169, 170, 188 Eip 28/29 gene, 244 Eliminius modestus, 161 Embryogenesis homologous structures, 16, 79 Crustacea, 68 Insecta, 23, 28,29, 31, 49, 55 Myriapoda, 57-8 juvenile hormone, 224 Endocrine physiology, eicosanoids, 185 Endogenous variation, 16 Endotoxin, 276

INDEX

see also Bacillus thuringiensis Engrailed genes, Arthropoda, 2, 8, 9, 15, 43 Engrailed proteins, Arthropoda, 57, 58, 68 Ensifera, 30 Ephemerella walderi, 141 Ephemeroptera, 141 Ephippigger ephippigger, 31 Epidermis homologous structures, 28 juvenile hormone, 226,246,249 imaginal discs and other imaginal precursors, 233-5 regulation of larval and pupal cuticle gene expression in higher Diptera, 229 regulation of larval cuticle gene expression, in Manduca, 226-9, 22 7 regulation of pupal and adult cuticle genes, 229-30 see also Pigmentation Epigenetic processes, Arthropoda, 79, 81 Epinephrine, 177 Epithelium, gut, 282-3, 284 Epoxidase, 121, 168 Epoxide hydrase, 217 1OR. 11S-Epoxybishomofarnesyl diazoacetate (EBDA), 248, 249 Epoxyeicosatrienoic acid, 117,121, 122 10,ll-Epoxyfarnesyl diazoacetate (EFDA), 247 Epoxygenase pathway, 168, 170, 186 Epoxyhomofarnesyl diazoacetate (EHDA), 246,248, 249 Escape behaviour, Arthropoda, 39-40, 67 Esculetin, 166,167, 170,170 Essential fatty acid, 117-9, 117, 118, 127-8, 180 see also Eicosanoids Esterase activity, 177, 178 Ethanolamine, 168 Ethrnostigmus rubripes, 57 ETYA (Eicosatetraynoic acid), 169,170, 188 Evaporative cooling, eicosanoids, 175-6 Even-skipped genes, 11, 15 Evolution, homology, 11 Excitatory neurons, 15

349

Excitatory postsynaptic current (EPSC), 325 Expression, glutamate receptors, 332-3 Eye, 76,234 Falck-Hillarp technique, 47 Farnesol, 244 Fascicle, 44 Fast extensor-tibiae (FETi), 24, 25 Fast motor neurons, Arthropoda, 15 Fat body eicosanoids, 117, 19CL7,191-3, 19.5 juvenile hormone, 216, 218, 232, 235 arylphorin, 235-7,236 JH-inducible larval proteins, 238 larval serum proteins in the Hemimetabola, 238-9 mechanism of action, 246, 247, 248. 249,250 methionine-rich storage proteins, 237 other JH-suppressible storage proteins, 237-8 Fatty acid, 117 see also Eicosanoids Fecundity, eicosanoids, 184 Feeding phase, juvenile hormone. 215, 218 Fenoxycarb, juvenile hormone, 214,254 Fertilization, eicosanoids, 198 Fevers, eicosanoids. 174-6 Fibre tracts, Arthropoda, 14 Fish, 160-1, 174 Flight eicosanoids, 132, 132, 172 homologous structures, 19, 20, 26-8. 27, 34-5, 64, 81 juvenile hormone, 216,235,239,241 Flightless grasshopper, 25 Fluid transport, eicosanoids, 184 FMRFamide, 23,40, 49-51, 59, 179 Fo$cula auricularia, 141 Forskolin, 332 Frogs, 174 Functional characterization, glutamate receptors, 332, 323, 334 G-neurone, Arthropoda, 36-7,38 G-proteins (Guanyl nucleotide binding protein), 1854,186, 223 GABA (Gamma aminobutyric acid), 23. 69, 330

350

Galleria, 237, 238 mellonella, 136, 138, 142. 187,229, 245 Gamma aminobutyric acid, 23, 69, 330 Ganglia, Arthropoda, 2, 7, 14 Gap junctions, gut, 282-3,283,2934, 295 Garter snakes, 161 Gas-chromatographic massspectrometric analysis (GC-MS), 135, 145, 172, 176 Gas-liquid chromatography, eicosanoids, 117, 129, 130, 149, 150 biosynthesis, 136, 137, 139, 142, 143, 144, 145 lipids, 133, 135 Genes Arthropoda, 7-10, 11, 12, 78-9 cascade, 11, 221-2 eicosanoids, 197 see also Epidermis Genitalia, juvenile hormone, 234 Geophilomorpha, 58 German cockroach, 140, 160, 238 Giant fibres, 46-7, 67-8 Giant interneurons, 39-40, 41 Gigaohm seal technique, 317-9, 320, 321,322 Glandular tissues, eicosanoids, 134 GLC see Gas-liquid chromatography Glomerular filtration rate, eicosanoids, 168 Glossina morsitans, 47 Glucagon, eicosanoids, 177 Glucocorticoid hormones, 197, 219 Glucose, glutamate receptors, 322 Glutamate receptors, 309-10, 334 central nervous system, 3 1 W , 311 cloning, expression and functional characteristics, 332-3 metabotropic, 331-2 visceral muscle, 330 see also Skeletal system Glutathione, 126, 130. 149, 195, 196 Glycoprotein, 77, 283, 289 Glycosaminoglycans, 282 GMP (Guanosine monophosphate), 223, 332 Goblet cells, gut, 282, 2834,283, 285, 286,289,292, 2934,295 Gomphocerus rufus, 219

INDEX

Gonad maturation, juvenile hormones, 219 Gonadotropin, 161 Gonyleptidae, 74 Granular phenoloxidase, juvenile hormone, 231-2 Granulocytes, 163 Growth, juvenile hormone, 215, 2 5 3 4 Gryllid, 25 Gryllus, 141 birnaculatus, 156 campestris, 35 GTP (Guanosine triphosphate), 223 Guanidinium, glutamate receptors, 323 Guanosine monophosphate, 223,332 Guanosine triphosphate, 223 Guanyl nucleotide binding protein (G protein), 18556,186, 223 Guanylyl cyclase, 223, 331 Gut Bacillus thuringiensis, 282-5,283 juvenile hormone 224,225 H-cell, 37 Haematin, eicosanoids, 195 Haematocytes, eicosanoids, 190, 196 Haeme, eicosanoids, 196 Haemocyctic immunity, eicosanoids, 162, 163 Haemocytes, eicosanoids, 134, 196 Haemoglobin, 195, 196, 238 Haemolin, eicosanoids, 162 Haemolymph eicosanoids immunity, 163, 164,164-6 lipid mobilization, 177-8 reproduction, 151, 152-3, 153, 154, 156 thermobiology, 175 juvenile hormone, 215, 216, 244, 24&7 epidermis, 230,234 fat body, 235, 238, 239 Hairs, Arthropoda, 67 Hallucinogen, 324 Harvestmen, 7 2 , 7 3 4 , 76 Heat shock protein, juvenile hormone, 244 Heliothis virescens, 290 zea, 129, 245 Helisoma duryi, 161

INDEX

Hematin. 149 Hemianax, 26 Hemiganglion, 17 Hemimetabola eicosanoids, 140 homologous structures, 17, 55, 56, 77 juvenile hormone, 216,235,238-9, 241 see also Schistocerca Hemiptera. 136, 141, 183, 214, 253 Hepoxilins, 117, 122, 123, 127 HETE see Hydroxypolyenoic fatty acid Heterodimers, 23, 220-1 Hetero-oligomers, 334 Heteroptera, 26, 82 High-performance liquid chromatography, eicosanoids, 117, 180 biosynthesis, 138, 144, 145 reproduction, 149, 151, 152, 153, 153 Hindgut, eicosanoids, 135 Hippodarnia convergens, 142 Histamine, 182 Histoblasts, juvenile hormone, 234-5 Holometabola eicosanoids, 140 homologous structures, 8, 17, 77 interneurons, 43, 52, 56 motoneurons, 25 juvenile hormones, 215, 216-7, 235, 239 see also Drosophila, Manduca, Tenebrio Hornarus, 64, 65, 66, 67 americanus, 174 Homeotic genes, Arthropoda, 78 Homo-y-linolenic acid, 118, 120, 121, 122,123,124,125, 167,176 biosynthesis, 136, 140, 142, 143, 144 lipids, 132, 132 oxygenation, 131 Homocysteate, 312, 333 Homodimers, steroid hormones, 220 Homology, nervous system, Arthropoda, 1-2,77-80, 82 concept of the identified neurone, 4-5 definition catalogue for, 14-16 criteria for, 12-14 historical aspects, 10-12 development and immunohistochemistrv. 7-10

351

morphology to genetics, 5-7 new approaches, 2-3 visual systems, 76-7 see also Chelicerata, Crustacea, Insecta, Myriapoda Homo-oligomers, 334 Homoptera, 140, 141 'Hormone response elements', 219 Hormones, definition, 219 ecdysteroid action, 221-2 eicosanoids, 185-6 peptide hormone action, 219, 222-3 steroid hormone action, 219-21,220, 223,248 thyroid hormone, 219, 220, 253 see also Juvenile hormone Host-parasite relationships, 180-1, 183 HPETE see Hydroperoxy fatty acids HPLC see High-performance liquid chromatography 5-HT (5-Hydroxytryptamine), 131, 179, 182 Humoral immunity, 162-3, 277 Hyalophora cecropia (silkmoth). 162. 226 Hydrogen Bacillus thuringiensis, 292, 293, 294, 295 gut, 283,283, 294,295 Hydroperoxidase, 195, 196 Hydroperoxy fatty acids (HPETEs), 119, 121, 122,125 5-HPETE, 122,125,126 12-HPETE, 179 Hydroperoxyendoperoxide, 122. 124 Hydroprene, 214,253 Hydroquinone, 149 2'-Hydroxy-4'-methoxyacetophenone. 183

2'-Hydroxy-4'-methoxypropiophenone1 183

20-Hydroxyecdysone, 225-6, 243, 244 epidermis, 227,228,232,233 fat body, 238 mechanism of action, 244, 250, 251, 252,253 muscle, 240 nervous system, 243 Hydroxyeicosatetraenoic acid, 117 Hvdroxvendoueroxide. 122.124

352

Hydroxypolyenoic fatty acids (HETEs), 119,121, 122,125 8-HETE, 161 12-HETE, 168 15-HETE, 154, 154, 180, 194, 195, 196 5-Hydroxytryptamine, 131, 179, 182 Hymenoptera, 142, 183 Hyperaemia, 182 Hyperpolarization, 312, 313 Hypersensitivity reactions, eicosanoids, 122, 162 Hypoenura, 141

INDEX

ventral nerve cord, 33-40,38, 41 motoneurons, 17-18 phylogenetic considerations, 24-8, 26,27 serial homologies, 18-24,21, 22 phylogeny, 80, 81, 82-3 segmentation, 79 sensory neurones interspecific homology, 31, 32-3 serial homology, 28-31,32 visual system, 77 Insecticides see Bacillus thuringiensis Insecticyanin, juvenile hormone, 227, 23G1 Insulin, 122, 185,223, 234 Interneurons glutamate receptors, 311 homologous structures, 7-8, 14, 16, 81 Chelicerata, 72 Crustacea, 67-8 Insecta, 19 immunoreactivity, 47-57,49,53-4, 56 suboesophageal ganglion, 36, 40-2, 42 supraoesophageal ganglion, 36, 42-7, 45 ventral nerve cord, 33-9,38,41 Myriapoda, 59, 61, 62 juvenile hormone, 242 Interphotoreceptor retinoid binding protein (IRBP), 249 Interspecific homology, Arthropoda, 31-3.32 Intracellular juvenile hormone receptors, 247 cytosolic binding proteins, 247-8 nuclear receptors, 248-50 Iodovinylmethoprenol, 249 Ion-selectivity, glutamate receptors, 323 Ion transport, eicosanoids, 16&73,170, 171,174, 184, 186, 186, 197-8 Iron protoporphyrin, 195 Isopoda, 68 Isoptera, 141, 142 Ixodes damimini, 182

Ibotenate, glutamate receptors, 311, 312,314,332,333 skeletal muscles, 315, 316. 323, 329-30 Ibuprofin, 180 Identified neurone, Arthropoda, 4 4 Idotea balthica, 63, 68 Imaginal discs, 215, 218, 225, 233-5 Immunity Bacillus thuringiensis, 277 eicosanoids, 122, 162-8, 1 6 6 7 , 175, 190, 196-7, 198 Immunocompetent cells, 163 Immunocytes, 163 Immunohistochemistry, Arthropoda, 6, 7-10, 33,46, 78 Immunomodulation, eicosanoids, 180-1 Immunoreactivity, Arthropoda, 47-8 crustacean cardioactive peptide, 51-4, 53,54 FMRFamide, 49-51 proctolin, 48-9, 49 serotonin, 54-7, 56 Indomethacin, 130, 157, 166,167, 170, 170, 171,171, 194, 195 Inflammation, eicosanoids, 122, 162 Information storage, glutamate receptors, 309 Inhibitors, eicosanoid biosynthesis, 183-4 Inhibitory neurons, 15, 24 Inositol triphosphate (IP3), 173,174,223 Insect growth regulators, 253-4 Insecta, neural systems, homology, 2 , 7 , 1617 compared to Crustacea, 68-9, 70 interneurons, 33 immunoreactivity, 47-57, 49,53, 54, Japanese beetle, 285 Juvenile hormone, 213-9,214,217, 56 254-5 suboesophageal ganglion, 36, 4G2, analogues as insect growth regulators, 42 supraoesophageal ganglion, 42-7, 45 253-4

INDEX

binding protein, 244, 246-7 embryonic actions, 224 esterase, 215,224 mechanism of action, 2446,245 binding proteins in the haemolymph, 246-7 intracellular receptors, 247-50,251 morphogenetic action, 251-3 premetamorphic actions muscle, 225,23941,240 other morphogenetic actions, 2 4 3 4 regulation of cellular commitment, 2254 receptor, 251-2 see also Epidermis, Fat body, Pigmentation Kainate, 312, 313, 314, 330, 332, 333 Kc cells, 243-4,245, 247 Kenyon cells, Arthropoda, 51 Ketamine, glutamate receptors, 324, 325 Keto-5,8,10,14-eicosatetraenoic acid (12KETE), 179 15-Keto-PGE2, 146, 154, 154 Ketoeicosatetraenoic acid, 117, 122 Kidney, 168-73,170,171,197 L-channels, glutamate receptors, 319, 320 Labopterella dimiditipes, 141 Labrum, Arthropoda, 43 Larval cuticle, juvenile hormone, 215 proteins, 227-9,227,236, 238, 250 Larval-specific storage protein, 239 LCP14 gene, 227-8,227,236,250 LCP14.6 gene, 227,228,236 LCP16/17 gene, 227,228-9,238,250 Lectins, 162, 313, 322-3 Legs homologous structures, 29, 30 juvenile hormone, 234 Lentil lectin, glutamate receptors, 322 Lepidoptera Bacillus thuringiensis, 276,277, 278, 2824,283,285,286 cell lysis, 292-4 mechanism of action, 288-9,295 binding protein, 246 eicosanoids, 128, 133, 1367,138, 142, 160,168

353

homologous structures, 26, 50,82 juvenile hormone, 214,215,216,217, 222,246 epidermis, 232,233,234 fat body, 236,236, 237 muscle, 239 premetamorphic action, 225 Lepisma, 16,21,25, 26,81,82 sacharina, 141 Leptinotarsa decemlineata (Colorado potato beetle), 237,238, 285 lineata, 50 Leucophaea maderae, 141, 198, 245,246 Leukocytes, 122 Leukotriene, 117, 122,126 A4, 122,126 B4, 180 C4, 126, 180 D4, 126 E4,126 Life histories, Arthropoda, 4 Life time, neuroblasts, Arthropoda, 18 Ligumia subrostrata, 169 Limulus, 77 polyphemus, 135, 175 Linden bug, 224 Linolenate, 117,118, 120, 128 . biosynthesis, 136,138, 139-40, 141, 142,142,143, 144, 145, 146 lipids, 132, 132 see also Dihono-y-linolenic acid, Homo-y-linolenic acid a-Linolenate, 118,120, 132, 132, 136-7, 138, 139, 144, 146 P-Linolenate, 118, 120, 132, 136, 144 Lipase, eicosanoids, 177, 178, 187 Lipids, eicosanoids, 116, 117, 1 3 1 4 , 132, 134, 167, 177-8, 177, 183, 187-8,189 Lipoxin, 121, 122,127 Lipoxygenase, 119, 121, 122,125, 126, 145, 194,195, 196, 197 fluid secretion rates, 168, 169-70,170 future discoveries, 185, 186, 188 immunity, 162, 166 neurophysiology, 179 reproduction, 154, 161 5-Lipoxygenase, 122, I25 Liquid filament switch technique, 329-30 Lithium, glutamate receptors, 323 Lithobiomorpha, 58

3 54

Lithobius, 26, 58-9, 60, 77. 83 fortificans, 61 Lizard, 174 Lobster, 55, 62, 67 Locusta decemlineata. 289 homologous structures, 8, 14, 18 compared to Crustacea. 64, 69, 70 interneurons, 35, 36, 39, 4&1, 49 motoneurons, 27 sensory neurons, 29, 31, 33 juvenile hormone, 224,239 migratoria eicosanoids, 158, 159, 159, 177, 187 glutamate receptors, 310, 311, 312, 314,3245,330 homologous structures, 36, 38. 40, 41, 50, 51, 52,53, 55 juvenile hormone, 216,245, 246 see also Schistocerca Lucilia sericata, 331 Lycoriella, 284 Lycosidae, 72 Lygaeus kalmii, 141 Lygus hesperus, 141 Lymantria dispar, 146 Lymnaea stagnalis, 161 Lysophospholipase, 188 Lysozymes, 162-3 Macrobrachium rosenburgii, 135 Macrophages, 122 Maleic acid, 167 Malpighian tubules, eicosanoids, 134, 135, 146, 197, 198 fluid secretion, 168-73, 170, 171, 176 future knowledge, 184, 185 reproduction, 150, 156 Mammals eicosanoids, 116, 119,121, 122-7, 123-7, 187, 188 fluid secretion rates, 168 immunity, 162 reproduction, 150, 156, 160 thermobiology, 174 glutamate receptors, 310 Mandibulata, 1, 2, 3, 6, 8, 12 compared Chelicerata, 69-70 interneurons, 4 4 , 4 6 , 5 1 4 , 5 5 phylogeny, 80 Manduca

INDEX

homologous structures, 17, 50 juvenile hormone binding protein, 246 embryonic actions, 224 intracellular hormone receptors, 24&9,250,251 mechanism of action, 222, 244 modulation of ecdysteroid action, 252,253 muscle, 23941,240 nervous system, 242-3 regulation of cellular commitment, 225-6 sexta eicosanoids, 134, 162, 163, 164, 165, 167, 187, 188-97,189,191-3,195 homologous structures, 50, 55, 56, 56,57 juvenile hormone, 216,217,218, 236,237,238, 245 see also Epidermis Mannose, glutamate receptors, 322 Mantid, 39-40 Markovian model, 318 Mass spectrometry, eicosanoids, 130, 133 Mast cells, 182 MDK (7s-methoprene diazoketone), 248,249, 250,2.51 Mead acid, 129 Meal beetle, 36, 37, 43, 52 Mechanoreceptors homologous structures, 29, 30, 62, 66. 67, 69 juvenile hormones, 242 Median prothoracic neurone, 45, 45 Megaohm seal technique, 317-8, 319-20, 321,322, 323, 324, 325, 330 Megoperculata, 73 Melanization, juvenile hormone, 231-3 Melanoplus bivattatus, 245 sanguinipes, 141 Membranes, eicosanoids, 136 Mesothoracic nervous system, Arthropoda, 19, 20.21, 22, 33, 34,37,38,79 Metabotropic glutamate receptors, 331-2 Metamery, Arthropoda, 7&9 Metamorphosis Arthropoda, 56 juvenile hormone, 213

INDEX

see also Premetamorphic actions Metathoracic nervous system, 19, 20,21, 22, 23, 33, 34-5, 37,38 Methionine-rich storage proteins, juvenile hormone, 237 Methoprene epidermis, 234 fat body, 237, 238 juvenile hormone, 214,2434,248, 249,253,254 muscle, 241 Methoprene-tolerant (Met), 247-8 Methyl anthranilate, eicosanoids, 183 Methyl farnesoate, juvenile hormone, 214,214 Methyl salicylate, eicosanoids, 183 Methyl transferase, juvenile hormone, 216 N-Methyl-D-aspartate (NMDA), glutamate receptors, 333, 334 central nervous system, 311, 312, 314 skeletal muscle, 317, 321, 330 4-Methyl-glutamic acids, 316 Methylsulphate, glutamate receptor, 330 Mexican grasshopper, 35 MHR3,250,252 Microdon albicomatus, 135 Microsomes, 178 Microvilli, gut, 282, 285 Midgut differentiation, juvenile hormone, 215 ‘Minuteness of resemblance’, Arthropoda, 13 Mitochondria1 proliferation, 241 Mitosis, 216 Modiolus demissus, 169, 185 Molecular biology Arthropoda, 3, 6, 11 eicosanoids, 197 glutamate receptors, 310 Molluscs, 161, 197 Monocytes, 122 Monophasic decay, 325 Monophylum, Arthropoda, 1-2, 57 Mosqui tos Bacillus thuringiensis, 284 eicosanoids, 132-3, 137, 14&7 Motoneurons glutamate receptors, 310-1, 312 homology, 7-8, 9, 14-15, 81, 83 Crustacea, 62-5, 63, 67 Insecta, 17-28,21,22, 26,27

355

Insecta and Crustacea compared, 68, 70 Myriapoda, 58-9, 60 juvenile hormones, 242, 243 Motor system, Arthropoda, 66, 72 Moulting see Ecdysis Mouthparts, Arthropoda, 29 Multiplicity of similarities, Arthropoda, 13 Multipolar neurones, 59 Musca domestica eicosanoids, 140, 145, 150. 159, 159 glutamate receptors, 314 homologous structures, 45, 47 Muscle eicosanoids, 198 glutamate receptors, 311 juvenile hormone, 225, 23941,240 nervous systems, 15, 59, 66, 81 Mushroom bodies, brain, Arthropoda, 2, 3, 81 Chelicerata, 69, 71, 74, 75 Insecta, 33, 43, 46, 51, 57 Myliostomata, 73 Myoglobin, 195, 196 Myriapoda, 2, 8, 10, 57-8, 82, 83 interneurons, 59, 61, 62 motoneurons, 58-9, 60 . segmentation, 79 visual systems, 77 Myristic acid, 118 Myrmica incompeta, 135 Myzus cerasi, 141 persicae, 139, 141 N-terminal Bacillus thuringiensis, 279.286.287-8 296,297 juvenile hormone, 247.249,255 steroid hormones, 218, 219 NAD (Nicotinamide adenine dinucleotide), 140, 157 NADP (Nicotinamide adenine dinucleotide phosphate), 140 Naiphoeta cinerea, 141 Naproxin, 170, 171, 194,195 Nauphoeta cinerea, 216,217 Navigational pathways, 29 Neck muscles, Arthropoda, 25-6 Nematocera, 284 Nernstian distribution, 284

356

Nerve roots, Arthropoda, 14 Nervous system, juvenile hormone, 225, 242-3 see also Homology Neuroamides, Arthropoda, 48 Neuroblasts, homology, 8-10, 12 Insecta, 18, 19,23,25,36, 41 Neuroendocrine, 169, 172-3 Neurogenesis, 3, 6, 8, 16, 79, 80 Crustacea, 68 Insecta, 18, 44, 48, 49 Myriapoda, 57-8, 59 Neurohormones, 64 Neuromodulator, Arthropoda, 48, 49 Neuropeptides, Arthropoda, 48 FMRFamide, 23, 40, 49-51, 59, 179 proctolin, 8-9,40,49, 64, 331 Neurophysiology, eicosanoids, 178-9 Neuropiles, Arachnida, 7, 7 3 4 , 75, 76 Neuropterans, eicosanoids, 133, 140, 142 Neurosecretion, 23 Neurotransmitters, 48, 49 New Zealand grass grub, 285 Nicotinamide adenine dinucleotide, 140, 157 Nicotinamide adenine dinucleotide phosphate, 140 Nicotinic acetylcholine receptors, 317 Nilaparvata lugens (brown rice planthopper), 184 NMDA see N-Methyl-D-aspartate Noctuids, 31 Non-spiking neurons, 15 Nuclear proteins, juvenile hormone, 229-30 29KDA, 250,251 Nuclear receptors, juvenile hormone, 248 larval epidermis, 248-50 hormonal regulation of, 250,251 Octocorals, 182 Octopamine, 265,331-2 Odonata, 19,26, 55,82, 141 Oedema, 162, 182 Oestrogen, 219,221 Oleicacid, 118,120, 132, 139, 140, 142, 143 1,2-0leoylacetylglycerol,332 Oligomerization, 298 Oligonucleotides, 249 Onchocerciasis, 278

INDEX

Oncopeltus fasciatus, 141, 216 Oniscus asellus, 55 Ontogeny, Arthropoda, 3-4,s-9, 12, 14, 29, 66 Onychophora, 2, 58, 71, 80 Oocyte maturation, 161, 218 Oogenesis, juvenile hormone, 218,224 Oothecins, 218 Opalescent glands, 159-60 Opilionid, 72,73, 74-6 Optic lobes, Arthropoda, 2, 44, 46,71, 75, 76 mass, Arthropoda, 74 nerves, Arthropoda, 74 Orconectes, 69 Organogenesis, 224 Orthopterans eicosanoids, 133, 140, 141, 144, 160 homology interneurons, 34, 3 5 4 , 39-40, 41, 46, 47, 54 sensory neurons, 30 Osmotic lysis hypothesis, Bacillus thuringiensis, 293-4 Osmotic protectants, 293 Ovaries, 147, 151, 153,153,246 Oviposition, 147,148, 154, 156-7, 159 Ovulation, 161 0x0-methylnorianoate, 140 Ozonolysis, 144, 145 Oxygenation, arachidonic acid, 129-31 Paeonol, 176 Palmitate (C16), 118, 132, 136, 142,143 Panaeus duorarum,l75 Pancreatic beta cell function, 185 Paper factor, juvenile hormone, 253 Papillia japonica, 285 Paragnath, Arthropoda, 66 Patch clamp studies, 313 Pathfinding, Arthropoda, 29, 31 Patinopecten yessoensis, 161 Pea lectin, glutamate receptors, 322 Penetrating tracheae, Arthropoda, 14 ‘Penknife model’, Bacillus thuriensis, 2968,297 Pentapeptide, Arthropoda, 48 Peptide hormones, 219,222-3 Peripatus, 2 Peripheral nervous system, Arthropoda, 31, 33

INDEX

Peripheral sensory system, Arthropoda, 81 Periplaneta, 26 americana eicosanoids, 134. 139, 141. 146, 178, 179,187 glutamate receptors, 312, 313, 314 homologous structures, 39, 40, 48, 49,49, 55, 56 juvenile hormone, 214,216 fuliginosa, 141 japonica, 141 Peritrophic membrane, gut, 283,288 Peroxidation, 119 Persistence, Bacillus thuringiensis, 278 Persistent storage protein, 239 Pesticides, 2534, 310 pH, gut, Bacillus thuringiensis, 276, 286-7, 286, 290, 292, 293-4, 298 Coleoptera, 28.5 Diptera, 284 Lepidoptera, 282, 285 Phalangiidae, 74 Phasmids, 2&8,27 Phencyclidine, glutamate receptors, 324-5 Phenoloxidase, 231. 232 Phenylalanine, 235 Philanthotoxin-343,326, 328, 329 Philanthotoxin-433 (PhTX-433), glutamate receptors, 324,326, 328, 329, 330 Philanthus triangulum, 327 Phosphatidylcholine, 117, 133-4, 151, 168. 187, 223 Phosphatidylethanolamine, 117, 133-4, 151, 168 Phosphatidylinositol 4,5-bisphosphate (PIP2), 117, 168, 223,246 Phosphatidylserine, 134 Phosphodiesterase, 131 Phospholipase, 223 A2, 117, 119, 151, 165,174, 187-8, 189 C, 173,174, 186, 188 Phospholipids Bacillus thuringiensis, 290 eicosanoids, 117, 119,121, 133-4, 135 biosynthesis, 136, 137, 138, 140, 144, 145, 146 fluid secretion rates, 174 immunity, 167 thermobiology, 176

357

Photoreceptors eicosanoids, 133-4 homologous structures, 2, 76-7 Phylogeny, Arthropoda, 3-4,6, 8, 11, 12. 13. 80-3. 82 Crustacea, 64, 68 Identified neurone, 5 Insecta, 24-8, 26,27, 33, 41, 45 Myriapoda, 57 segmentation, 78 Pieris brassicae, 129, 146 Pigmentation, juvenile hormone, 2 15, 216,225,230 cuticular melanization, 231 dopa decarboxylase, 232-3,233 granular phenoloxidase, 231-2 insecticyanin, 23&1 Planococcus citri, 141 Plasmatocytes, 163 Platelet aggregation, 182, 184 Plecoptera, 141 Pleuropods, Arthropoda, 29 Plexaura homomella, 182 Plurisegmental interneurons, Arthropoda, 33 Polyamine amides, glutamate receptors, 326,327-9 Polyphagan beetles, 50 Polyphylum, Arthropoda, 2 Polyspermic fertilization, 161 Polyunsaturated fatty acid, 116, 117, 128-9 C18, 119,120, 129, 132-3 C20, 116, 119,120, 129, 131-6, 132. 134

docosahexaenoate, 118, 132, 146 eicosanoid biosynthesis, 119-222, 120, 121 essential fatty acids, 117-9, 118, 127-8 see also Arachidonic acid, Biosynthesis, Eicosanoids. Eicosapentaeonate. Homo-ylinolenic acid, Linolenate, Oleic acid, Palmitate, Stearate Pore formation, Bacillus thuringiensis, 291, 2968,295,297 Position, criterion of, Arthropoda, 13 Potassium Bacillus thuringiensis, 282, 2834, 283, 285, 286, 289, 291, 2924,295 eicosanoids. 168, 179 glutamate receptors, 323

INDEX

358

Praying mantis, 39, 41 Precis coenia, 2 2 5 4 Predator avoidance, prostaglandins, 182-3 Premetamorphic actions, juvenile hormone, 2154,217 muscle, 23941, 240 nervous system, 242-3 other morphogenetic actions, 243-4 regulation of cellular commitment, 225-6 see also Epidermis, Fat body, Pigmentation Principle proleg retractor muscle (PPRM), 240,240, 242 Procambarus, 26, 65, 68 clarkii, 55, 62 Procian yellow, 20 Prociphilis fraxinifolly , 141 Proctolin, 8-9,40,49, 64, 331 Prolactin, 253 Proleg retractor muscles, 239-41,240, 242 Prolegs, juvenile hormone, 23941,240 Prophenoloxidase, 163 Proprioceptors. 29, 30, 66 Prostacyclin, 124, 182 Prostaglandin A*, 123, 182, 185 fluid secretion rates, 169, 170, 173 predator avoidance, 182 reproduction, 153, 153, 154, 154, 160 tobacco hornworm tissues, 191,191-3, 192-3, 194,19.5 Prostaglandin B2, 123, 154, 154, 160, 194 Prostaglandin D2, 123, 124 blood flukes, 180 reproduction, 154, 154 thermobiology, 176 tobacco hornworm tissue, 191. 191-3, 193,194,197 Prostaglandin dehydrogenase, 156, 157, 158, 182 Prostaglandin E?, 123, 124, 129-31, 144, 145, 198 ecological significance, 180, 182, 184 fluid secretion, 168, 169, 172, 173 lipid mobilization, 177-8, 177 neurophysiology, 179 reproduction, 147, 150, 153, 153, 154, 154,155, 1569, 160 thermobiology, 174, 175, 176 tobacco hornworm. 191.191-3. 193. 194

Prostaglandin F2, 123, 124, 130, 144, 145 ecological significance, 182 fluid secretion rate, 172 neurophysiology, 179 reproduction, 149, 150, 153, 153, 154, 154, 157, 158, 159, 160, 161 thermobiology, 176 tobacco hornworm, 191, 191-3, 192, 193, 194 Prostaglandin G , 124, 195 Prostaglandin H, 124, 194, 195, 196, 197 Prostaglandin I, 124, 154, 154 Prostaglandins, 116, 117, 119, 121, 122, 123,124 arachidonic acid oxidation, 129-31 biosynthesis, 144, 145, 146 ecological significance, 1 8 0 4 fluid secretion, 168-73, 170, 171 lipid mobilization, 177-8, 177, 183 neurophysiology, 178-9 reproduction, 14741,148, 153, 154, 1.55, 159 thermobiology, 174-6 Prostate gland, 160, 198 Proteases. gut, 285,287 Protein kinase A , 223 Protein kinase C, 223, 332 Proteins eicosanoids, 191, 191 homology, 13, 68 juvenile hormone, 216, 219. 222, 226, 2274,227,235,241 see atso Cry proteins, Cyt proteins, Fat body Prothoracic systems, Arthropoda, 19, 22,33,34 Prothoracicotropic hormone (PTTH), 215,218 Protocerebral bridge, 69,74 Protocerebrum, Arthropoda, 43, 44, 45, 4.5, 46, 56 ‘Proton peril’ hypothesis, Bacillus thuringiensis, 292-3 Protoxins, Bacillus thuringiensis, 287, 288,292 Pterothorax, 25 Pterygotes, 30, 40, 81 PUFAs see Polyunsaturated fatty acid Puffing response, juvenile hormones. 243 Pupal gin traps. 240, 242-3 Pyriproxyfen: 214, 234, 239, 254 .

I

INDEX

Pyrrhocoridae, 253 Pyrrhocoris apterus, 224 Quisqualate, 332, 333, 334 central nervous system, 311, 312, 314 metabotropic, 331 skeletal muscle, 315-6, 330 see also Channel gating kinetics Quisqualis indica, 315

Radiation, Arthropoda, 4, 5, 12 Radioimmunoassay, 117 Radioligand binding studies, 185 Rana escutenta, 168 temporaria, 168 Receptor organs, Crustacea, 65-7 Receptor proteins, eicosanoids, 187 Receptors Bacillus thuringiensis , 288-90 juvenile hormone, 254,255 see also Intracellular juvenile hormone receptors Rectum, 169, 184, 198 Reproduction eicosanoids, 130-1, 135, 147-61, 148, 153,154,155, 159, 184, 198 juvenile hormone, 213, 218,219 Reptiles, prostaglandins, 160-1 Resistance, Bacillus thuringiensis, 278, 290 Reticulitermes Jovipes, 142 Retinas, eicosanoids, 133-4 Rhodnius. 224 prolixus, 55, 183-4, 216,218,245, 246 Rhodopsin, 249 Ribonucleic acid glutamate receptors, 332-3 juvenile hormone, 228-30, 232,246, 250,251,255 fat body, 2367,236, 238 Rilaena triangularis, 73, 7 4 6 , 75, 76 Riptortus clavatus, 214, 239 RNA see Ribonucleic acid Rostrostomata, 73 Rubidium, glutamate receptors, 323 s-channels, glutamate receptors, 319, 320

359

S1 cells, Arthropoda, 55 Salicaceae, 183 Salicylaldehyde, 183 Saltatoria, 26, 82 Salticidae, 72 Sarcophaga, 331 bullata, 47,55, 56, 229, 234, 235, 2365,245 Scarabaeid grubs, 285 Sceloporus jarrovi, 161 Schistocerca homologous structures, 79,83 motoneurons, 17, 19, 20. 21,21, 22, 24, 64 sensory neurons, 31,32 americana, 55 gregaria glutamate receptors, 312, 313, 314. 315-6.319-28, 329-30 homologous structures, 35, 36, 40, 55 nitens, 49 Schistosoma mansoni, 180-1, 181 Sclerotization, juvenile hormone. 232 Scorpions, 72 Secretion rate, fluids, eicosanoids, 168-72, 170,171 Segmentation, Arthropoda, 33, 78-9 Semaphore concept, Arthropoda, 8 Seminal fluids, 160, 198 Sense organs, Arthropoda, 66, 8 1 Sensilla basiconica. 28 companiformia, 28 trichoidea, 28 Sensory neurons homology, 39, 66. 83 interspecific homology, 31, 33 serial homology, 28-31,32 Juvenile hormone, 242-3 Septate junctions, gut, 282,283 Serial homology, Arthropoda. 14, 18-19, 28-3 1.32 other motoneuron types, 23-4 typical motoneurons, 19-23.21, 24 Serotonin immunoreactive intraganglionic neurones, 75-6 Chelicerata, 72, 73 Crustacea, 64 Insecta, 37,40, 42, 44, 45, 45. 54-5,56 Myriapoda, 59 Serratia marcescens, 163, 164

360

Sesquiterpenoid, juvenile hormone, 213 Setae, juvenile hormone, 215 Sex pheromone, juvenile hormones, 218-9 Sex specificity, storage proteins, 237 Shore crab, 51 Shrimp, 65 Silicic acid column chromatography, 151, 158 Silk glands, juvenile hormone, 225 Silkmoth, 162, 226 Simple eye, Arthropoda, 77 Simuliid blackfly, 278, 284 Sipyloidea sipylus, 27 Skeletal system, glutamate receptors, 310,312,313,314-5, 331 developmental studies, 329 ibotenate-sensitive receptors, extrajunctional membrane, 329-30 see also Channel gating kinetics SKF-525A, 170,170 Smooth muscles, eicosanoids, 198 Sodium Bacillus thuringiensis, 282, 284, 285, 291 eicosanoids, 168, 196 glutamate receptors, 323 Solubility, Bacillus thuringiensis, 286-7 Soma, Arthropoda, 14 Southern corn rootworm, 285 Soybean lectin, glutamate receptors, 322 Species, homology, 12-13 Specific quality, criterion of, Arthropoda, 13 Specificity, Bacillus thuringiensis, 278 Spermathecae, 149, 150, 151-3, 153, 160 Spermatophore, eicosanoids, 131, 134, 147, 149, 151, 152 Spiking neurons, 15 Spiking transmission, 69 Spirobolus marginatus, 83 Spodromantis linealoa, 35 Staining methods, Arthropoda, 17-18, 20 Starfish, 161 State switching phenomenon, 317 Stearate (C18), 118, 132, 136, 137,138, 142,143, 144, 146, 147 Steroid hormones, 219-21,220, 223, 248 Stomatogastric system, 43 Storage proteins see also Fat body

INDEX

Stretch receptors, Arthropoda, 30 Suboesophageal ganglion, Arthropods, 36, 4G2,42,45, 72, 73-4, 75 Succinyl-concanavalin A, 322, 323 Supraoesophageal ganglion, Arthropoda, 36, 42-7, 45, 68, 70, 74-6, 75, 76 Surface patterning, juvenile hormone, 225 Sweating, eicosanoids, 176 Symbiosis, eicosanoids, 139 Symploce capitata, 141 Synapomorphy, 5 , 7 , 11, 12, 80 Synaptic physiology, eicosanoids, 198 Systematics, 8, 11

Teleogryllus commodus eicosanoids, 134,134, 172, 198 biosynthesis, 141, 142, 143, 144-5 reproduction, 147-53, 154,155, 158, 159 glutamate receptors, 314, 316, 329, 331-2 juvenile hormones, 216 oceanicus, 241 Tenebrio, 43, 50-1, 52 molitor eicosanoids, 134, 135, 142, 146, 159, 172 homologous structures, 36, 40-1, 42, 45, 47, 49, 49, 52,53,54, 55, 56, 56, 57 juvenile hormone, 230 Testes, 135, 149, 152, 158, 160 Tetrachloroisoquinoline, 327 Tetramethylammonium, glutamate receptors, 323 Tettigoniida, 30, 31, 33 Thermobia domestica, 188 Thermobiology, eicosanoids cicadas, 175-6 mediation of behavioural fevers, 174-5 Thin-layer chromatography, 117, 130, 139, 144, 187, 190, 194, 196 ecological significance, 182 immunity, 168 reproduction, 149 thermobiology, 176 Thoracic nerves, Arthropoda, 17, 18, 19-23,21, 22, 24, 44, 45, 69 Thorax, 135

INDEX

361

Thromboxane, 117,119,121,124, 154, 154,

183-4

Thyroid hormone, 219,220,253 Thysanura, 39,141 Tibicen, 176 dealbatus, 135 Ticks, 178 Tipulid leatherjacket, 284 TLC see Thin-layer chromatography Tobacco hornworm see Manduca sexta Tracheata, 57-8,71,81,83 Tracheoblasts, 241 Triacylglycerols, 133 Triatoma infestans, 159,159 Triboliuin, 8 castaneum (red flour beetle), 183 Trichoplusia ni, 129,142,158,159,159,

237,238 2,4,6-Trihydroxyacetophenone,183 10,11,12-Trihydroxy-5,8,14,17eicosatetraenoic acid, 161 Trimetaphan, glutamate receptors, 324, 325,327 Trioxilin, 117, 122,123,127 Trisaccharide raffinose, 293 Tritocerebral commissure giant interneuron, 35-6 Tritocerebrum, Arthropoda, 43,45,45,

56 Tubocurarine, glutamate receptors, 324 Tumour growth, eicosanoids, 122 Tympani, Arthropoda, 30,31,33,34 Type, organism, 10 Tyrosine, 223,235 Ultraspiracle protein, 221 ‘Umbrella model’, Bacillus thuriensis,

297,298 Urine. 169

compared to suboesophageal ganglion, 42 compared to supraoesophageal ganglion, 43,44,45 immunoreactivity, 49, 50, 52,54-5,

54,57 Myriapoda, 59,61 Ventral unpaired median neuron, 18 Vertebrata, 14,309-10 Visceral muscles, glutamate receptors,

330,331 Visual ganglia, 76 Visual systems, Arthropoda, 7&7 Vitamin E, 117 Vitellogenesis, juvenile hormone, 216,

218,221,246 VNC see Ventral nerve cord Voltage clamp experiments, Baciflus thuringiensis, 292 VUM (Ventral unpaired median neuron), 18 Water transport, 168,197 Waxmoths, 136-7,146 Wheatgerm lectin, glutamate receptors,

322 Wing malformations, essential fatty acids, 128 Wing stretch receptor, 16, 19,30,31 .Wingless genes, Arthropoda, 8,9, 43 Wings, juvenile hormone, 234 World Health Organization, 278 Xenopus, 313,333 laevis, 332 Xiphosura, 71-2 X-ray crystal structure, Bacillus thuringiensis, 276,279,280,281,

296 Yellow fever mosquito, 134,169,170,

V-ATPase, gut, 284,292,294,295 Vasodilation, eicosanoids, 162 Ventral external oblique muscle (VEO),

172

Zinc, steroid hormones, 219,220 Zootermopsis angusticollis, 140,141 Ventral nerve cord, Arthropoda, 2,6,7, Zophobus, 26 81 morio, 40 Insecta, 17,3340,38,41 Zygentoma, 26

240,241


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