Advances in Insect Physiology
Volume 31
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Advances in Insect Physiology edited by S. J. Simpson Department of Zoology and University Museum of Natural History, University of Oxford, Oxford, UK
Volume 31
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Contents Contributors
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Foreword
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Obituary
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Insect Sterol Nutrition and Physiology: A Global Overview SPENCER T. BEHMER, W. DAVID NES
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The Nutritional Physiology of Aphids ANGELA E. DOUGLAS
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The Neurobiology of Taste in Insects STEPHEN M. ROGERS, PHILIP L. NEWLAND
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Trehalose – The Insect ‘Blood’ Sugar S. NELSON THOMPSON
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INDEX
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Contributors S. T. Behmer Department of Zoology, University of Oxford, South Parks Road, Oxford, OX1 3PS, UK A. E. Douglas Department of Biology, University of York, PO Box 373, York, YO10 5YW, UK P. L. Newland School of Biological Sciences, University of Southampton, Bassett Crescent East, Southampton, SO16 7PX, UK W. D. Nes Department of Chemistry and Biochemistry, Texas Tech University, Box 41061, Lubbock, TX 79409-1061, USA S. M. Rogers Department of Zoology, University of Cambridge, Downing St, Cambridge CB2 3EJ, UK S. N. Thompson Analytical Chemistry Instrumentation Facility and Department of Entomology, University of California, Riverside, CA 92521, USA
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Foreword This is my first volume as Editor and I must begin by thanking, on all our behalves, Peter Evans for the excellent job he has done over the past 17 years. Peter has overseen the publication of 11 volumes of Advances in Insect Physiology, each one an adventure. I hope to live up to the very high standards he has set. The flavour of a journal will inevitably reflect the tastes of its editor, as well as the prevailing trends and advances in the subject. Recent years have seen a growing interest in physiological mechanisms among ecologists, behavioural scientists and evolutionary biologists. We should nurture this trend, and I aim actively to commission papers in these areas. I am also keen to receive unsolicited proposals, submissions and requests for papers in any area of insect physiology – and I am willing to interpret the field broadly. As the present volume was nearing completion, the sad news came that Reg Chapman had died. Reg’s book The Insects: Structure and Function (editions 1–4) has been the pre-eminent text and reference work on insect physiology for the past 34 years, and will continue to be so for some time yet. Through this seminal book Reg has had a greater role in disseminating the subject than any other insect physiologist. I would like to dedicate Volume 31 to Reg. This is especially fitting, given the direct influence he had through his research on the topics covered by the reviews in this volume. As a result, although it was not planned as such, this volume serves as a Festschrift for Reg.
Reginald Frederick Chapman (1930–2003): An Obituary After taking his undergraduate degree in zoology at Queen Mary College, University of London, in 1951, Reg accepted a scholarship from the AntiLocust Research Centre to work on the factors controlling roosting behaviour in locusts. The ALRC was based at the Natural History Museum in London. Its director was B.P. Uvarov, the father of locust biology. He had brought together a distinguished research team that included John Kennedy and Donald Gunn; two others who, along with Uvarov, had a formative influence on Reg’s scientific development. Reg’s doctoral research was carried out at Birkbeck College in London, to where he would later return and write The Insects. After his Ph.D., Reg joined the International Red Locust Control Service and travelled to East Africa, where he lived and worked for more than three years in the Rukwa Valley, a locust outbreak area in a remote, largely uninhabited, area in East Africa, with no electricity, telephone or radio. Reg built his own house, complete with plumbing, grew his own fruit and vegetables, and became addicted to the idea that laboratory and fieldwork should be combined. At first he extended his study of roosting behaviour in locusts (as well as studying frogs and lizards), then he turned to feeding behaviour – the area in which he went on to make his most important scientific contributions. An indication of Reg’s dedication to taking the laboratory to the field is seen in his experiments testing Peggy Ellis and Graham Hoyle’s hypothesis that haemolymph potassium levels influenced locust behaviour through a direct effect on neuromuscular transmission. This required running a flame photometer under the most unpromising of circumstances. Reg left the locust service in 1957 and took a lectureship at the University of Ghana, where he worked on tsetse flies and grasshoppers, before moving in 1959 to a position at Birkbeck College in London. There he taught invertebrate zoology and developed a masters course in entomology, which he taught single-handedly. Writing and giving the more than 100 lectures and associated practical classes in the course provided Reg with an encyclopaedic knowledge of entomology, and also honed his extraordinary teaching skills. He brought together this knowledge in The Insects – Structure and Function, first published
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in 1969. While at Birkbeck, Reg was promoted from Lecturer to Reader, then Professor. In addition to his formidable teaching schedule, he established a productive and innovative research group, focusing on feeding behaviour and the chemosensory system. It was during this time that Liz Bernays, who was to become Reg’s wife, joined the group, as did another key figure, Wally Blaney. In 1970, Reg moved to become Director of the Laboratory Research Division of the Anti-Locust Research Centre (later to be renamed the Centre for Overseas Pest Research), which had moved to new premises in Wright’s Lane, Kensington. The incoming Director, Peter Haskell, gave Reg the freedom to develop research projects both in the lab and overseas in the field. Reg and his team took as their themes the basis of varietal resistance in crop plants and the mechanisms of host–plant selection by phytophagous insects. He directed and played an active, practical part in research projects that included a study of movement patterns in the brown rice planthopper, based in the Philippines, a project in India investigating varietal resistance of sorghum to insect pests, one in Botswana on sorghum entomology, another in Nigeria on a pest grasshopper, the impact of soil termites on grasslands, and the impact of pesticide residues on non-target species, and a radar-tracking study in Mali on grasshopper migrations. Meanwhile, in the laboratory, Reg, Liz and their colleagues and students continued to explore feeding behaviour and host–plant selection, making significant progress in elucidating the role of host–plant chemistry. In 1983, Reg and Liz left for Berkeley – Liz to a Faculty position and Reg to an unsalaried situation. In so doing, he had chosen to put Liz’s career ahead of his own, which was a typical example of Reg’s belief in supporting talented others above himself. In 1987, the University of Arizona took advantage of the opportunity to get Chapman and Bernays and, to Berkeley’s loss, offered them both professorships. Reg was appointed Professor in Insect Neurobiology in John Hildebrand’s Division of Neurobiology. Until his retirement in 2001, Reg played an influential role in the academic and teaching life at Tucson, which, thanks in no small part to his efforts, has become one of the world’s most distinguished centres for insect science. One especially notable achievement was Reg’s contribution to the establishment and success of the crossdepartmental Center for Insect Science, for which he served as Interim Director from 1998–1999. Reg also continued his research on feeding and chemosensory physiology, wrote numerous scientific papers, co-wrote the book Host–Plant Selection by Phytophagous Insects (1994) with Liz, co-edited and part-wrote Regulatory Mechanisms in Insect Feeding (1995), and completely rewrote and reillustrated The Insects (1998), as well as writing several key review papers and assuming a demanding graduate and undergraduate teaching load. He did all of this and more under the shadow of advancing leukaemia, which had first been diagnosed in 1990 and to which he finally succumbed on May 2nd 2003.
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Reg’s legacy lies at several levels. Through The Insects – Structure and Function and its numerous translations, he has reached and influenced many thousands of biologists worldwide. In his other, more specialist books, book chapters, and review papers, Reg has shaped thinking in the field of insect–plant relationships through his ability to bring together large amounts of scattered literature and build new syntheses. This was especially well demonstrated in his classic review in Advances in Insect Physiology (1982), in which he collated vast amounts of published data on chemoreceptor numbers in insects (itself a huge job) and used them to build functional arguments to explain variation in chemoreceptor numbers between different taxa and lifehistories. At the time such a ‘comparative approach’ was virtually unknown, but has since become a cornerstone of modern biology. Reg also published 110 primary research papers, typified by their scholarship and attention to detail. Many more papers that could have borne his name were published under the authorship of his 37 doctoral students, and the many others to whom he gave his time and encouragement freely. Reg’s philosophy for supervising graduates was to ‘‘try to convey three things: integrity, knowledge and enthusiasm. . . This requires developing a level of mutual trust with each individual that enables us to assess work critically’’. Those of us fortunate enough to have been supervised by Reg have seen his integrity, knowledge and enthusiasm first hand – as well as his passion for getting it right and his ability to grasp the point and effortlessly spot the flaws in apparently water-tight arguments. To this list of attributes, those of us who knew Reg would also wish to add his modesty, kindness, inspiration – and the twinkle in his eye. Reg is survived by two children from his first marriage, Anne and Philip, two grandchildren, and his beloved wife, Liz Bernays. Steve Simpson, June 2003
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Insect Sterol Nutrition and Physiology: A Global Overview Spencer T. Behmera and W. David Nesb a
Department of Zoology, University of Oxford, South Parks Road, Oxford OX1 3PS, UK; b Department of Chemistry and Biochemistry, Texas Tech University, Box 41061, Lubbock, TX 79409-1061, USA
1 Introduction 2 2 Sterol structures and dietary sources 4 2.1 Sterol structure and nomenclature 4 2.2 Algal and lichen sterols 4 2.3 Fungal sterols 6 2.4 Plant sterols 7 2.5 Animal sterols 9 3 Insect sterol use and metabolism 9 3.1 Insect sterol use 12 3.2 Insect sterol metabolism 20 3.3 Considerations of patterns of sterol use and metabolism 35 4 Insect sterol physiology 38 4.1 Sterol taste and the regulation of intake 42 4.2 Sterol absorption 44 4.3 Sterol transport and tissue distribution 47 4.4 Sterol reproductive physiology 52 5 Insect sterol ecology 54 6 Applied implications and evolution of sterol metabolic constraints 56 7 Conclusions 57 Acknowledgements 59 References 59
Abstract Unlike most animals, insects lack the capacity to synthesize sterols that are required in lipid biostructures, as precursors to important steroid hormones and as regulators of developmental processes. Therefore insects must acquire sterols from their diet. Hundreds of different sterols have been identified and the review starts by documenting the occurrence of sterols in different insect foods. Next we look at the various nutritional and biochemical studies that have been conducted, and organize them according to insect relatedness, which allows insect sterol use and metabolic capabilities to be viewed from an evolutionary perspective. How sterol structure influences insect feeding behavior is ADVANCES IN INSECT PHYSIOLOGY VOL. 31 ISBN 0-12-024231-1 DOI: 10.1016/S0065-2806(03)31001-X
Copyright # 2003 Elsevier Ltd All rights of reproduction in any form reserved
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examined, and the fate of sterols once they have been ingested, including the processes of absorption and transport, their distribution in different tissues, and their role in reproduction, is detailed. The extent to which sterols may influence ecological outcomes is also considered, especially in phytophagous insects with known sterol metabolic constraints. Finally, mention is made of the potential use of exploiting insect sterol requirements and constraints for pest control, as well as the ability of insects to adapt to the presence of novel sterols in their diet. 1
Introduction
The dietary need for sterols in insects was first established in the blowfly, Lucilia sericata (Hobson, 1935a), and is the only proven nutritional difference between insects and most other animals. However, it was the seminal work by Clark and Bloch (1959a) that demonstrated the inability of insects to synthesize sterols de novo, thereby generating sterol auxotrophy. Since these early studies, the dietary requirement for sterols has been extended to other insect orders, including Orthoptera, Blattaria, Hemiptera, Coleoptera, Diptera, Lepidoptera, and Hymenoptera (Svoboda et al., 1994). Initial studies on insect sterol nutrition focused on cholesterol and its surrogates. For instance, Clayton (1964) reared the hide beetle, Dermestes vulpinus, on a diet that contained different amounts of cholesterol and cholestanol supplemented in the diet and determined that cholesterol played two roles in governing larval growth and development. The two dietary sterols differ in the structure of the nucleus, i.e., the existence of a 5- (double bond at carbon 5–6) or 0- (no double bond), respectively, but this difference has only a minimal effect on the amphipathic properties of the molecules. This means that both can serve equally well as a membrane insert. Clark and Bloch (1959a) surmised that cholestanol, added as a bulk sterol to the food, was serving a structural role, whereas cholesterol added at trace levels acted as a sparing sterol, functioning in a metabolic role. At that time the exact nature of the metabolic role was speculative. Further study demonstrated that ecdysteroids derived from cholesterol were functioning hormonally to control insect growth and maturation (reviewed by Gilbert et al., 2002). The importance of sterol availability and structure to the growth of phytophagous insects was examined in subsequent years by a research group at the U.S. Department of Agriculture (Thompson, 1984). In a beautifully designed set of pioneering experiments, they opened-up the area of structure– activity and enzymatic studies, including the rational design and assay of inhibitors of insect sterol metabolism. For example, certain vertebrate hypocholesterolemic agents such as triparanol were found to inhibit important enzymatic reactions in the sterol biosynthetic pathway that were involved in the conversion of phytosterols to cholesterol in the tobacco hornworm, Manduca sexta, and other insects (Svoboda and Robbins, 1967). These agents effectively stopped insect growth and development, and thus became important tools for the study
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of phytosterol metabolism in other insects (Svoboda, 1984). Following inhibitor treatment the identification of desmosterol (cholesta-5,24-dienol) as the terminal intermediate to cholesterol in M. sexta provided the first information on intermediate steps involved in the pathway. Subsequently, Ritter and Nes (1981a) composed a sterol-free diet, which was used to culture Helicoverpa zea. The availability of this diet proved instrumental in establishing the exactness of sterol homeostasis (balance in the type and amount of sterol) in controlling insect physiology. At about the same time, Kircher (1982) at the University of Arizona was studying sterol ecology, and Ikekawa et al. (1993) in Japan was studying the stereochemical mechanisms involved in phytosterol side chain dealkylation and reduction. The latter investigators demonstrated the relationship of side chain transformations in insects to more derived animal systems. More recently, the effect of sterols on the Hedgehog (Hb) gene family, which encodes a group of secreted signal molecules that are essential for growth and patterning of many different body parts (Ingham et al., 1991; Alexandra et al., 1999), has been documented. Beachy and coworkers have found that cholesterol can modify covalently the Hedgehog family proteins in animals, thereby affecting morphology (Porter et al., 1996). These observations open up exciting new avenues of research regarding the multiple roles of sterols in insects distinct from studying the role of cholesterol as a membrane insert or as a precursor to ecdysteroids. The role of sterols in insect nutrition and physiology (Clayton, 1964; Dadd, 1977, 1985; Svoboda and Thompson, 1985; Bernays, 1992), including sterol metabolism and ecdysteroid production (Robbins et al., 1971; Thompson et al., 1973; Svoboda et al., 1978; Svoboda and Thompson, 1985; Svoboda and Feldlaufer, 1991; Ikekawa et al., 1993; Svoboda, 1999), has been documented regularly over the past 30 years. Our goal in this chapter is to provide a global overview of this topic, but for practical reasons we have deliberately downplayed the details of sterol metabolism and ecdysteroid production since they have been presented elsewhere on numerous occasions. We have three specific aims. Our first is to look for unifying themes that can explain patterns of sterol use and metabolism among the insects that have been studied to date. We do this by examining sterol use and metabolism within an evolutionary framework, paying particular attention to insect phylogeny. Next, we review different aspects of sterol physiology by following the journey of sterols through the body. Here we discuss a range of topics, including whether insects taste sterols and regulate their intake, the processes related to sterol metabolism, absorption, transport, and tissue distribution, plus the role of sterols in reproduction. Our third aim is to explore the manner in which insect sterol nutrition and physiology can impact ecological and evolutionary processes. Related to this is the potential of targeting insect sterol metabolic limitations for pest management in agricultural systems. Finally, we conclude the chapter by discussing how the unique sterol physiology of insects might be extended to new and developing areas of sterol research.
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Sterol structures and dietary sources
Insects are incredibly diverse, with approximately one million species so far described. Estimates including undescribed species suggest that the true number could range between 2 and 5 million (Speight et al., 1999), and Erwin (1982) predicts this number is much higher. The unmatched diversity of insects as a group is partly attributed to their ability to use a wide range of substrates as food, including plants, fungi, algae/mosses, animal tissues and decaying organic materials. As detailed in the following sections, sterols exist in numerous forms but there are some underlying features in the types of structures distributed in the food supply of insects. 2.1
STEROL STRUCTURE AND NOMENCLATURE
Sterols normally ingested by insects are characterized by three domains: a 3 hydroxyl group (A), a tetracyclic ring system (B), and a side chain of 8–10 carbon atoms (C), as shown in Fig. 1. The polar and non-polar elements, consisting of domains A versus B and C, give rise to the amphipathic nature of the molecule. This feature, when coupled with the side chain oriented to the ‘‘right’’, provides a suitable fit of the sterol into the lipid leaflet of membranes (Nes et al., 1978; Bitmann, 1997). The sterol may possess methyl groups at C-4 and they are defined accordingly as C-4 dimethyl sterol, C-4 monomethyl sterol and C-4 desmethyl (no methyls) sterols. The degree of C-4 methylation affects the hydrogen bonding ability of the C-3 hydroxyl group as evidenced in the thin-layer chromatography of these compounds (Xu et al., 1988). Alternatively, the sterol may possess a C-24 methyl or ethyl group, which may possess stereochemistry as either - (in front) or - (in back) oriented. It should be noted that the stereochemical nomenclature for the side chain C-24 alkyl groups can be affected by neighbouring substituents when the R/S-nomenclature is used and that the configuration assignments can be different for groups in the side chain relative to the nucleus (Parker and Nes, 1992). The prefix in -sitosterol is dropped in common usage for chemical reasons (Nes and McKean, 1977; Thompson, 1984). The structure assignments for sterol used typically by phytosterol biochemists is different from natural product chemists, who follow the recently revised numbering of sterols by the International Union for Pure and Applied Chemistry (IUPAC) and International Union for Biochemistry (IUB), as indicated in Fig. 1. In the current chapter, we continue to use the sterol nomenclature preferred by traditionalists. 2.2
ALGAL AND LICHEN STEROLS
Feeding on algae and lichens (a symbiotic relationship between algae and fungi) is relatively rare among the insects, but it is found among the Plecoptera, Orthoptera and Psocoptera and, to a small degree, in some of the Hemiptera
INSECT STEROL NUTRITION AND PHYSIOLOGY
FIG. 1 Sterol numbering systems. A, B and C represent structural domains.
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and Endopterygotes. Most aquatic insects include algae in their diets, but few specialize on it. Some insects, including tree-dwelling Collembola and some Plecoptera, feed on a mixture of algae and lichens. In contrast, the lichen grasshopper, Trimerotropis saxatilis (Orthoptera: Acrididae) and the lichen-eating caterpillar, Manulea replana (Lepidoptera: Acrtiidae), are lichen specialists. Perhaps not surprisingly, they have also become highly cryptic on their food. Several hundred species of algae have been examined for sterol content, and the types of sterols found are usually related to taxonomic position (Patterson, 1994). In general, all the algae, except for the dinoflagellates (Pyrrophyta), have predominantly 5-sterols. Among the freshwater algae that might be eaten by insects, the Charophyta (stoneworts) have two dominant sterols, 24ethylcholesterol and isofucosterol, while the major sterols in Chrysophyta (golden algae) and Xanthophyta (yellow-green algae) are 24-ethylcholesta5,22-dienol and 24-ethylcholesterol, respectively. Cryptophyta (cryptomonads) are characterized by the presence of 24-methylcholesta-5,22-dienol, while the Chlorophyta (green algae) contain 24-epi sitosterol (clionosterol) and ergosterol, plus a few other sterols in smaller quantities (Seckbach et al., 1993). Typically, orders with multicellular algae are characterized as having 24-ethylcholesterol and isofucosterol as their primary sterols, while the more primitive, primarily unicellular orders have a range of sterols, including compounds with double bonds at the 5-, 7-, and 5,7-position. Within the algae, ergosterol has so far only been found in unicellular chlorophyte species belonging to the orders Volvocales and Chlorococcales, and in a few members of the Euglenophyta (Patterson, 1994; Volkman, 2003). A variety of sterols have been identified in the Cyanobacteria (blue-green algae), and they frequently have an ethyl group at C-24 and a ring system with 5-, 7-, or 5,7-double bonds (Nes and Nes, 1980; Rzama et al., 1994). 2.3
FUNGAL STEROLS
Fungus eating is primarily found in 6 insect orders (Collembola, Zoraptera, Psocoptera, Thysanoptera, Coleoptera and Diptera), although individuals in 3 additional orders also occupy this feeding guild (Thysanoptera, Lepidoptera and Hymenoptera). Perhaps the best known fungus-feeders are the leaf cutting ants, which maintain fungus gardens within their nests. Leaf-cutting ants (Hymenoptera: Formicidae: Myrmicinae) include a taxonomically compact group of 12 genera that comprise the tribe Attini and contains approximately 190 species (Weber, 1972). Their distribution is confined to the Nearctic and Neotropical biogeographic region, and the fungus-growing Macrotermitinae seem to be their closest ecological equivalent (Wood and Thomas, 1973). Ambrosia beetles are another well-known example of a fungus-eating insect. The fungi are composed of three major groups, the Zygomycota (bread moulds), Ascomycota (sac fungi) and Basidiomycota (club fungi), and well over a hundred species have been examined for sterol composition (Weete, 1973;
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Patterson, 1994). In general, ergosterol is the dominant sterol, although 5and 7-sterols are occasionally found, and C28 sterols are much more frequent than C29 sterols (Weete, 1973). Ergosterol and related 5,7-sterols are the dominant sterols in most of the Zygomycota, but related C28 sterols can also be abundant and some species contain cholesterol and 24-methylcholesterol (Beilby and Kidby, 1980) or 24-ethylcholesterol (Beilby, 1980). Ergosterol is also the principle sterol in most Ascomycota, but some orders, such as Tuberales and Tafrinales, often contain brassicasterol in large proportions (Weete et al., 1985; van Eijk and Roeymans, 1982). Among the Basidiomycota, ergosterol and related C28 sterols tend to be the rule, although sterols with a 7-monoene nucleus have been recorded in this group (Weete and Laseter, 1974). Other fungal sterols include fungisterol, which is found in bracket fungus (Fomes applanatus), and zymosterol, a sterol biosynthesis intermediate is found in some yeast species (Myant, 1981). In the Ascomycota, Gibberella fujikaroi synthesizes as many as 35 sterols (Nes et al., 1988a) and the formation of 24 -methylsterols (ergosterol) and 24 -methylcholesterol is known to occur via different post-lanosterol pathways (Nes and Lee, 1990). 2.4
PLANT STEROLS
Herbivory is found in only 9 of the 29 insect orders (Collembola, Orthoptera, Phasmida, Thysanoptera, Hemiptera, Coleoptera, Lepidoptera, Diptera and Hymenoptera), but almost half of all described insect species occupy this feeding niche (Strong et al., 1984). Phytophagous insects can be found on every plant species and are known to feed on a range of plant tissues and parts, including leaves, stems, wood, roots, buds, flowers, fruit, pollen, nectar, xylem and phloem. At least 100 different sterols have been identified in plants so far (Akihisa et al., 1991), with structural variation occurring mainly in the position and extent of nuclear and side chain unsaturation and in the extent of 24-alkylation in the side chain. With respect to sterol side chain differences, the size of the 24-alkyl group and its direction ( - or -oriented) reflect phylogenetic differences in plant evolution, with the ergosterol side chain (6)
SCHEME 1 Hypothetical pathway to plant sterols.
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found in primitive organisms and the stigmasterol side chain found in derivedorganisms (5) (Scheme 1) (Nes and McKean, 1977). Bryophytes are primitive land plants that contain primarily 5-sterols, although some species have been found to contain 5,7-sterols (Nes and Nes, 1980). With respect to the orientation of the alkyl group at C-24, bryophytes contain both and forms, which contrasts with the fungi and algae where the orientation is usually only (Patterson, 1994). Among the pteridophytes, a group of seedless, vascular plants consisting of whisk ferns, horsetails and clubmosses, sitosterol is the dominant sterol but campesterol and stigmasterol are often present (Chiu et al., 1988). These authors also found that in the pteridophytes, sitosterol and stigmasterol have their C-24-ethyl groups in the orientation. With respect to campesterol, they found that in lycopods it tended to follow the orientation, but in the more derived horsetails, whisk ferns and true ferns it followed the orientation. Among the conifers that have been examined, 5-sterols are common, with sitosterol being the most dominant (Svoboda et al., 1995b; Schiff and Feldlaufer, 1996). Flowering plants (Class Angiospermae) are by far the largest group of vascular plants on earth today, with approximately 250 000 identified species representing a relatively small number of families (320–418) (reviewed by Heywood, 1993). The 5-sterols sitosterol (24 ) and stigmasterol (24 ) are the most common and abundant sterols found in the angiosperms, but many species also contain campesterol (24 -methyl) and 24-dihydrobassicaterol (24 -methyl), usually in a 2:1 ratio (Patterson, 1994). Angiosperms, in contrast to the more primitive plants, sometimes contain 7-sterols, such as spinasterol (24 ) and stigmast-7-enol (24 ) (Nes and Nes, 1980). The order Caryophyllales is particularly rich in 7-sterols with 7 of the 12 recognized families having major and sometimes dominant quantities, and lesser quantities of 5- and 0sterols (Salt et al., 1991). For example, a survey of the family Curcurbitaceae revealed that it contained almost exclusively 7-sterols (Akihisa et al., 1987). Sterols with a 7 configuration have also been reported in individual members belonging to the Leguminosae, Theaceae and Sapotaceae (Bergmann, 1957). However, sterols with a 5,7 nucleus, such as that found in ergosterol, have yet to be found in large amounts in any angiosperms. In broad terms, most plants contain multiple sterols that vary within some general theme, and there seems to be an underlying phylogenetic pattern determining sterol nucleus structure. An extreme case of structural variation within a species can be found with corn, where 24-alkyl olefin formation is characteristic, but as many as 60 different sterols can be synthesized (Guo et al., 1995). Many insects, especially bees, include pollen in their diets. In general the pollen for most plant species accumulate intermediates such as 24methylenesterols and 9 ,19-cyclopropyl sterols (Nes and Schmidt, 1988; Lusby et al., 1993). Occasionally, however, sitosterol is the principle sterol, as Standifer et al. (1968) found in mule fat (Baccharis viminea), juniper (Juniperus uthaensis), heartsease (Polygonum sp.), waterleaf (Hydrophyllum
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capitatum) Scotch pine (Pinus sylvestris), European alder (Alnus glutinosa) and Lombardy poplar (Populus nigra var. italica). These same authors also found stigmasterol in large amounts in the pollen of heather (Calluna vulgaris) and cholesterol in the pollen of both cottonwood (Populus fremontii) and cat’s ear (Hypochaeris radicata). They found little evidence, however, that taxonomy could be used to predict pollen sterol profile since three species belonging to the family Salicaceae varied widely in the content of C27, C28 and C29 sterols. 2.5
ANIMAL STEROLS
Zoophagous and saprophagous lifestyles are found in all insect orders, with the exceptions of the Phasmida and Zoraptera (Southwood, 1973), and although the range of animals that insects will eat is quite broad, vertebrates and arthropods (including other insects) seem to constitute the bulk of an insect’s diet. In the vertebrates, cholesterol is by far the most abundant sterol, and the majority of it persists in free (unesterified) form in the plasma membranes, the subcellular membranes of cells and in myelin of nervous tissue (Myant, 1981). Additionally, all vertebrates appear capable of synthesizing cholesterol, but in general they do not absorb C-24 alkylated sterols from their diets (Nes and McKean, 1977). The phylum Arthropoda is the largest of the animal kingdom and sterol composition and metabolism have been extensively studied in this group. It is generally accepted that arthropods cannot synthesize sterols and that they acquire sterols from their foods. Cholesterol tends to be the principle sterol in most species, especially in carnivores and scavengers, but there are numerous exceptions, particularly among those that eat plants and/or fungi (Nes and McKean, 1977; see Section 3.2). Among the phytophagous insects, cholesterol tends to be the dominant sterol, although phytosterols are also often present, and sometimes in large quantities (see Section 3.2). 3
Insect sterol use and metabolism
To date slightly more than 100 insects have been investigated with respect to sterol use or metabolism (Table 1). However, no systematic examination has been made with regard to sterol use/metabolism, and of those insects that have been studied, half come from the Coleoptera, Lepidoptera and Hymenoptera. Among the remaining species to have been examined, only 8 orders are represented. An additional bias exists in that many of the species studied are agricultural pests. In spite of these shortcomings, we document what is known about sterol use and metabolism in different insects and look for emerging patterns, paying particular attention to evolutionary relationships among insects (both within and between orders) and to their differing natural histories. Throughout the paper we used the Tree of Life (Maddison et al., 2001), and to a lesser extent Borror et al. (1989), to determine the relatedness
10
TABLE 1 Survey of sterol studies among the different insect orders. Diet composition taken from Southwood (1973). Order
Number of unique species
Utilization studies (no. of species)
Profile studies (no. of species)
Algae/Lichen/Decaying Plant Material Plant Products Algae/Detritus Animal Algae/Lichen/Animal Dead Plant Material/Lichen/Moss Plant Plant/Animal/Mixed
0 1 0 0 0 0 0 9
– – – – – – – 9
– 1 – – – – – 2
Animal Fungus/Mites Mixed/Wood/Animal Decaying Plant Material Moss/Dead Insects Algae/Fungus/Dead Plant Material Animal Plant
0 0 3 0 0 0 0 0
– – 1 – – – – –
– – 2 – – – – –
S. T. BEHMER AND W. D. NES
Archaeognatha (bristletails) Thysanura (silverfish and firebrats) Ephemeroptera (mayflies) Odonata (dragonflies and damselflies) Plecoptera (stoneflies) Embiidina (web-spinners) Phasmida (stick or leaf insects) Orthoptera (crickets, locusts, grasshoppers and katydids) Mantophasmatodea Zoraptera Dictyoptera (cockroaches, termite and mantids) Dermaptera (earwigs) Grylloblattodea (ice bugs and rock crawlers) Psocoptera (barklice, booklice) Phthiraptera (lice) Thysanoptera (thrips)
Diet
Total
Plant
1
–
1
Plant
1
–
1
Plant/Animal/Mixed Animal Animal Animal Plant/Animal/Mixed Animal
9 0 0 1 21 0
– – – – 18 –
9 – – 1 4 –
Plant/Fungus/Animal Plant/Fungus/Animal Animal Animal Animal/Mixed Plant
2 10 0 1 0 18
2 10 – 1 – 16
– 2 – – – 4
Plant Plant/Animal/Mixed
8 20
– 2
8 18
105
60
53
INSECT STEROL NUTRITION AND PHYSIOLOGY
Hemiptera (sucking bugs) Sternorrhyncha (psyllids, whiteflies, aphids and coccids) Auchenorrhyncha (cicadas, leafhoppers, treehoppers and fulgoroids) Heteroptera (true bugs) Megaloptera (dobsonflies and alderflies) Raphidioptera (snakeflies) Neuroptera (lacewings, antlions and owlflies) Coleoptera (beetles) Strepsiptera Diptera (flies) Nematocera (long-horned flies) Brachycera (short-horned flies) Mecoptera (scorpion-flies) Siphonaptera (fleas) Trichoptera (caddisflies) Lepidoptera (moths and butterflies) Hymenoptera (sawflies, wasps, bees and ants) Symphyta (sawflies) Apocrita (bees, wasps and ants)
11
12
S. T. BEHMER AND W. D. NES
of different insect families to one another, and have organized the studies according to insect order ! superfamily ! family ! genus ! species. 3.1
INSECT STEROL USE
Traditionally sterol use has been explored by rearing insects on artificial diets that contain different sterols. This approach requires, however, that an artificial diet already exists or is readily formulated, and that the diet does not have a large pool of contaminating sterol. For example, Behmer and Grebenok (1998) have shown that basic ingredients like wheat germ contain sitosterol, while casein and albumin contain small amounts of cholesterol. Likewise, Nes (1987) demonstrated that agar from red algae contains cholesterol, agar from corn contains a mixture of phytosterols and cholesterol and that Tween 80 and some commercial phospholipids contain a mixture of phystosterols. In such instances contaminating sterols need to be removed, via chemical extraction, or it needs to be shown that they have negligible effects. Demonstrating the latter can prove difficult, however, because of the sparing effect associated with cholesterol (e.g. Clark and Bloch, 1959a). For this particular reason, some insect sterol utilization studies should be viewed with caution, especially those in which the purity of sterols and diet ingredients is not reported. Nonetheless, artificial diet studies have revealed some strong trends, the most notable being that cholesterol supports normal growth and development in all predaceous insects, and most herbivorous insects. Sterol use in phytophagous insects can also be studied by manipulating plant sterol profiles via fungicides and transgenic techniques. In general, though, sterol use studies are lacking for most insect orders, and among the Exopterygota, only grasshoppers have been extensively studied. 3.1.1
The Exopterygota
Grasshoppers are one of the few groups of insects in which sterol use has been systematically studied, with the eight selected species coming from two families, the Romaleidae and Acrididae, and within the latter group four different subfamilies being represented. Inspection of the results from these various species indicates a robust pattern in sterol use among grasshoppers. Cholesterol and sitosterol, two 5-sterols, plus cholestanol, a 0-sterol lacking an alkyl side chain, were found to support growth better than the other sterols examined. Additionally, cholesterol, sitosterol and cholestanol were the only sterols that supported growth from hatchling to the adult stage (Dadd, 1960; Behmer and Elias, 1999a, 2000). Most of the grasshoppers that have been studied, with the exception of the grass specialist, Locusta migratoria, are generalist feeders (i.e. they eat mixed plant diets). This means that they may regularly ingest plant material that contains unsuitable sterols (e.g. 5,22, 7 and 7,22). To investigate the consequences of this on growth and development, Behmer and Elias (1999a, 2000) reared Schistocerca americana hatchlings on synthetic diets
INSECT STEROL NUTRITION AND PHYSIOLOGY
13
containing different amounts and proportions of suitable and unsuitable sterols. The results were somewhat surprising because grasshoppers failed to complete development on diets when the ratio of suitable to unsuitable sterols in the food fell below 70%. Interestingly development was impaired even when the suitable sterol was present in amounts that alone would have supported growth. Results from these experiments indicate that in grasshoppers the cholesterol sparing mechanism is weak and that their tolerance for the intake of unsuitable sterols is low. Sterol use in grasshoppers has also been examined by using plants with modified sterol profiles. For instance, Charlet et al. (1988) applied the systemic fungicide fenpropimorph to wheat, which caused a change in the sterol profile from mostly 5-sterols, e.g. sitosterol, to a mixture of 8-sterols and 9 ,19cyclopropylsterols (95% of the total). They found that newly hatched L. migratoria nymphs reared on the treated wheat took longer to develop relative to nymphs reared on control wheat, and that nymphs on the treated wheat exhibited wing reversals and reached the adult stage after four instars, rather than the normal five. Chemical analyses revealed that locusts reared on the modified wheat had markedly different tissue sterol profiles, and reduced titre of ecdysteroids, compared to locusts reared on normal wheat. The only other exopterygotes reared on diets containing different sterols are the omnivorous cricket Gryllulus domesticus (Chauvin, 1949) and the cockroach Blattella germanica (Noland, 1954; Gordon, 1959). Like grasshoppers, both grew equally well on diets containing cholesterol and sitosterol. In contrast, however, crickets also grew well on diets containing ergosterol and moderately well on diets containing stigmasterol, while cockroaches grew well on desmosterol and ergostanol, but not zymosterol. 3.1.2
The Endopterygota
Also called the Holometabola because of the dramatic changes that occur between the larval, pupal and adult stages, four of the five largest insect orders (Coleoptera, Diptera, Lepidoptera and Hymenoptera) can be found in this group. Many agricultural pests are endopterygotes and the development of artificial diets for mass rearing purposes has greatly facilitated sterol studies in these insects. A relatively large number of beetles have been examined with respect to sterol use but they all come from one suborder, the Polyphaga. It is, however, the largest suborder within the Coleoptera and contains approximately 300 000 described species, which are split into16 superfamilies. To date, representative species from five of the superfamilies within Polyphaga have been examined for sterol use (Bostrichoidea, Cucujoidea, Tenebrionoidea, Chrysomeloidea, and Curculionoidea). Overall, there seems to be a remarkable flexibility among beetles to use a relatively wide range of sterols, including ones with 5-, 5,22-, and 5,7,22-bonds (Table 2). There is, however, one notable exception. It would
14 TABLE 2 Survey of sterol use among the different insect orders. Insects are organized first by order and then by family. When more than one family occurs within an order, shared roman numerals indicate they come from the same superfamily. No inferences are made about the relatedness of superfamilies to one another. The key to the table is as follows: ( þ þ þ ) ¼ strong growth; ( þ þ ) ¼ moderate growth; ( þ ) ¼ weak growth; (0) ¼ no growth and ( ) ¼ not studied. Numbers in parentheses indicate the cited study. Zoo sterols
Species Cholesterol
Dictyoptera (cockroaches, termites and mantids) Blattellidae Blattella germanica (5, 6) Coleoptera (beetles) Dermestidae (I) Dermestes vulpinus (7, 8, 9, 10, 11) Attagenus piceus (12) Trogoderma granarium (13, 14) Anobiidae (I) Lasioderma serricorne (8) Stegobium paniceum (8)
7-Dehydrocholesterol
Cholestanol
0 0
þþþ þþþ
Sitosterol
Other sterol (1)
Other sterol (2)
Stigmasterol
0 0 0 0 0 0
spinasterol 0 0 0 0 0 0
lathosterol
þ þ þ þ þ þ
þþþ þþþ
0 0
0 0
0 0
þþþ
þþ
þ þ þ þ þ þ
þ þ þ þ þ þ
Other sterol (3)
þþþ
0
þþ
þþþ
þþþ
þþþ þþþ þþþ
0 þþ þþþ
0 0 þþþ
0
0
þþþ
þþþ
þþþ þþþ
þþþ þþþ
þþ þþ
þþþ þþþ
lanosterol 0 0
Fungal sterols Ergosterol
Zymosterol
0 0
0 0
0 0 0 0
þþþ
desmosterol þþþ
ergostanol þþ
0
0 0
0
þþþ þþþ
þ 0
S. T. BEHMER AND W. D. NES
Orthoptera (locusts, grasshoppers, crickets and katydids) Ensifera (short-horned grasshoppers) Acrididae (I) Locusta migratoria (1) þþþ Schistocerca gregaria (1) þþþ Schistocerca americana (2,3) þþþ Trimerotropis pallidipennis (2) þþþ Barytettix humphreysii (2) þþþ Melanoplus differentialis (2) þþþ Romaleidae (I) Romalea guttata (2) þþþ Taeniopoda eques (2) þþþ Caelifera (long-horned grasshoppers) Gryllidae (II) Gryllus domesticus (4) þþþ
Plant sterols
Diptera (flies) Nematocera (long-horned flies) Culicidae Aedes aegypti (29) Culex pipiens (30) Brachycera (short-horned flies) Drosophilidae (I) Drosophila melanogaster (31, 32) Drosophila pachea (33) Calliphoridae (II) Lucilia sericata (34, 35) Phormia regina (36) Calliphora erythrocephala (37) Cochliomyia hominivorax (38)
þþþ
þþþ
þþ
þþ
þþþ
þþþ
þþ
þþþ
þþþ þþþ
þþþ
þþ
þþþ
þþþ þ þþþ
þþþ þþþ
þþ 0 þ þ þ þ
þ þ þ þ
þþþ
þþþ
þ þþþ
þþþ þ
þ þ þ þ
þ þ þ þ
þ þ þ þ
lanosterol 0
þþ
þþ
þþþ þþþ
þ
0 þþþ þþþ
lanosterol 0 0
þþþ
þþþ
þþ þþ
þþ þ
þþþ þþþ
þþþ þþþ
þ þþþ
þ
þþþ 0
þþþ 0
0
þ
þ
þþþ
0
þ þþþ
þ þ þ þ
þþþ
desmosterol þþþ
þþþ
þþþ þþþ þ þþþ þþþ þþþ
þþþ þþþ
campesterol þþþ
þþþ
þþþ þþþ
desmosterol
campesterol
fucosterol
þþþ
þ
þþþ
desmosterol 0
schottenol
lathosterol
þþþ
þþþ
þþ þþþ
INSECT STEROL NUTRITION AND PHYSIOLOGY
Ptinidae (I) Ptinus tectus (8) Silvanidae (II) Oryzaephilus surinamensis (8) Tenebrionidae (III) Tribolium confusum (15, 16) Tenebrio molitor (17) Cerambycidae (IV) Hylotrupes bajulus (18, 19) Bruchidae (IV) Callosobruchus chinensis (20) Acanthoscelides obtectus (21) Curculionidae (V) Hylobius pales (22, 23) Anthonomus grandis (24) Sitophilus granarius (w/o symbionts) (25) Sitophilus oryzae (w/ symbionts) (25) Scolytidae (V) Xyleborus ferrugineus (26, 27) Scolytus multistriatus (28)
þ
þþþ 0
þþ þþþ
þ þþ
0
0
0
(Continued )
15
16
TABLE 2 Continued Zoo sterols
Species Cholesterol
Sarcophagidae (II) Pseudosarcophaga affinis (39) Sarcophaga bullata (40) Muscidae (III) Musca domestica (41, 42, 43, 44, 45, 46) Anthomyiidae (III) Delia brassicae (47) Siphonaptera (fleas) Xenopsylla cheopis (48)
þþþ
Other sterol (1)
Other sterol (2)
Other sterol (3)
Ergosterol Zymosterol
7-Dehydro- Cholestanol Sitosterol Stigmasterol cholesterol
þþþ þ
þ þ
þþþ þþ
þ
þ þ
Fungal sterols
campesterol þ
desmosterol þ
þ þþþ
þ
þþþ
þþ þþþ
þþ
þþþ
þ
þþþ
þ
þþþ
þ
þþþ þþþ
þþþ þþ
þþþ þþ
þþþ
þþþ
þþþ þ
þþþ þþþ þþþ
þ
þþ þþþ
þþþ þþþ
þþþ
0
spinasterol þþ
þþþ 0 þþþ þ þ þ þ
þ þ þ þ
þ þ þ þ
0 þþþ þ þþþ
þþþ
þ
þ þþþ
0
þþ þþþ
þþþ
campesterol þþþ
brassicasterol þ
fucosterol þþþþ
0
S. T. BEHMER AND W. D. NES
Lepidoptera (moths and butterflies) Plutellidae (I) Plutella xylostella (49) Gelechiidae (II) Pectinophora gossypiella (50) Sitotroga cerealella (51) Tineidae (III) Tineola bisselliella (52) Tortricidae (IV) Homona coffearia (53) Argyrotaenia velutina (54) Pyralidae (V) Ectomyelois ceratoniae (55) Anagasta kuehniella (56) Pyrausta nubilalis (57) Corcyra cephalonica (58) Crambidae (V) Crambus trisecta (52) Diatraea grandiosella (59) Bombycidae (VI) Bombyx mori (60, 61) Sphingidae (VI) Manduca sexta (62)
þþþ þþþ
Plant sterols
Hymenoptera (sawflies, wasps, bees and ants) Apocrita (distinct waist) Chalcididae (I) Brachymeria lasus (65) Pteromalidae (I) Pachycrepoideus vindemiae (65) Apidae (II) Apis mellifera (66)
þþþ þþþ
0
þþ
þþþ
þþþ
þ
þþ
þþ
þþþ
þþ
þþ
þþ
þþþ
þþ
þþþ
þþ
spinasterol þþ
lathoterol þ
brassicasterol þ
þ
campesterol 24-methylenecholesterol þ þþþ
(1) Dadd, 1960; (2) Behmer and Elias, 1999a; (3) Behmer and Elias, 2000; (4) Chauvin, 1949; (5) Noland, 1954; (6) Gordon, 1959; (7) Fraenkel et al., 1941; (8) Fraenkel and Blewett, 1943; (9) Levinson, 1962; (10) Bergmann and Levinson, 1966; (11) Budowski et al., 1967; (12) McKennis, 1947; (13) Agarwal, 1970; (14) Sehgal and Agarwal, 1973 (15) Fraenkel and Blewett, 1943; (16) Magis, 1954; (17) Leclercq, 1948; (18) Rasmussen, 1956; (19) Rasmussen, 1958; (20) Ishii, 1951; (21) Chiu and McKay, 1939; (22) Clark, 1973; (23) Richmond and Thomas, 1975; (24) Vanderzant, 1963; (25) Baker, 1974; (26) Norris et al., 1969; (27) Chu et al., 1970; (28) Galford, 1972; (29) Goldberg and De Meillon, 1948; (30) Dadd and Kleinjan, 1984; (31) Cooke and Sang, 1970; (32) Cooke and Sang, 1972; (33) Heed and Kircher, 1965; (34) Hobson, 1935a; (35) Hobson, 1935b; (36) Brust and Fraenkel, 1955; (37) Sedee, 1961; (38) Gingrich, 1964; (39) House, 1954; (40) Goodfellow et al., 1971; (41) Monroe et al., 1961; (42) Kaplanis et al., 1965; (43) Dutky et al., 1967; (44) Silverman and Levinson, 1954; (45) Levinson and Bergmann, 1957; (46) Bergmann and Levinson, 1966; (47) Dambre-Raes, 1976; (48) Pausch and Fraenkel, 1966; (49) Behmer and Grebenok, 1998; (50) Vanderzant and Reiser, 1956; (51) Chippendale, 1971; (52) Fraenkel and Blewett, 1946; (53) Sivapalan and Gnanapragasam, 1979; (54) Rock, 1969; (55) Levinson and Gothilf, 1955; (56) Fraenkel and Blewett, 1943; (57) Beck et al., 1949; (58) Sarma and Sreenivasaya, 1941; (59) Chippendale and Reddy, 1972a; (60) Ito, 1961; (61) Ito et al., 1964; (62) Svoboda and Robbins, 1968; (63) Ishii and Urushibara, 1954; (64) Ritter and Nes, 1981; (65) Thompson, 1981; (66) Herbert et al., 1980.
INSECT STEROL NUTRITION AND PHYSIOLOGY
Arctiidae (VII) Chilo simplex (63) Noctuidae (VII) Helicoverpa zea (64)
17
18
S. T. BEHMER AND W. D. NES
appear that dermestid beetles have a limited ability to use fungal sterols, which is in contrast to the two other beetle families, e.g. anobiids and ptinids, within the superfamily Bostrichoidea, which grow quite well on the fungal sterol ergosterol. Perhaps sterol use in the Bostrichoidea is related to feeding ecology. For the zoophagous D. maculatus (Dermestidae) cholesterol is the only sterol that supports growth and development, while in the phytophagous Kharpa beetle, Trogoderma granarium, which is also a dermestid, sitosterol and stigmasterol, in addition to cholesterol, support growth and development. Among the anobiid larvae, the ability to grow and develop on diets containing ergosterol is probably related to the fact that they are often found living in fungi. It would be interesting to know whether these different sterol utilization abilities follow any phylogenetic pattern (cholesterol only ! cholesterol þ phytosterols ! cholesterol þ phytosterols þ fungal sterols). The other interesting finding among the beetles is that lanosterol, a 8,24cholesterol analogue, does not support growth and development in the two species in which it was tested (Norris et al., 1969; Chu et al., 1970; Clark, 1973; Richmond and Thomas, 1975). The Diptera are the second largest insect order and are divided into two suborders, the Nematocera and Brachycera. Within the Nematocera, which generally include small insects with delicate long antennae, 16 superfamilies exist but to date sterol use has only been examined in two species, and both these come from the mosquito family, Culicidae. The larvae of most mosquito species feed on algae and organic debris, but a few are predaceous and feed on other mosquito larvae. Sterol use in Aedes aegypti (Goldberg and De Meillon, 1948) and Culex pipiens (Dadd and Kleinjan, 1984) is broad, although C. pipiens only grows on non-cholesterol diets when lecithin is present. The Brachycera, which include the more compact flies with short antennae, are divided into 20 superfamilies. Sterol use, however, has only been examined in species from four of these superfamilies, and all of these come from the section Calyptra`tae, within the division Schizo´phora. Inspection of the data in Table 2 reveals that sterol use in this group of insects is highly variable. The first notable difference occurs within the Drosophilidae, whose larvae tend to feed on fungi found in decaying fruits and vegetables. Drosophila pachea, which breeds in the stems of senita cactus, is perhaps the most famous example of unique sterol use among the insects. Heed and Kircher (1965) found that 7-stigmasten-3 -ol, isolated from the cactus or synthesized, but not cholesterol, sitosterol, stigmasterol or, surprisingly ergosterol, could replace the cactus in the diet of flies reared non-aseptically or axenically. In contrast, D. melanogaster, which is considered a generalist feeder, grows well on diets containing this latter group of sterols (Cooke and Sang, 1970, 1972). The cabbage root fly, Delia brassicae, is a specialist feeder on plants in the family Brassicaceae. Like other species in the Anthomyiidae, it feeds on plant roots, an environment that favours fungal growth, and perhaps its sensitivity to
INSECT STEROL NUTRITION AND PHYSIOLOGY
19
cholesterol (Dambre-Raes, 1976) suggests it has evolved and become a sterol specialist much like D. pachea. Among the muscids only Musca domestica has been examined, and it seems that a range of sterols can be used, although of those tested cholesterol is clearly superior (Table 2). Perhaps this reflects the generalist feeding nature of its larvae, which are found feeding on excrement and various types of decaying material. Flies within the superfamily Oestro`idea, which includes the Calliphoridae and Sarcophagidae, tend to feed on carrion, excrement and similar decaying material, so it is not surprising that cholesterol supports strong growth for flies in this group. It is interesting, however, that the plant sterol sitosterol supports growth and development to high levels in three species (Table 2). This may reflect the remarkable ability of some insects to spare cholesterol while using other sterols for structural purposes (e.g. Clark and Bloch, 1959a). An additional finding of interest for flies from these two families, especially considering the environments in which these insects feed, is that the utilization of fungal sterols seems to be poor or even non-existent. The Lepidoptera are another large order and the majority of the larvae are phytophagous, although a small number are predaceous and some are ectoparasitic. There are five suborders, but all of the species that have so far been studied for sterol use come from the suborder Ditrysia. Of the 17 superfamilies in this suborder, seven are represented in Table 2. The most notable trend is that sitosterol supports strong growth and development in all the species for which it has been tested, which is not surprising considering it is the most common and typically abundant phytosterol. Stigmasterol (5,22) is also widespread in plants, although usually present in smaller quantities than sitosterol, and is also readily used by most lepidopteran species. There are exceptions, however, and specialists that feed on plants where stigmasterol tends to be absent or rare, e.g. Plutella xylostella (crucifers), Pyrausta nubilalis (grasses) and Bombyx mori (mulberry), seem to utilize it less well relative to sitosterol. Performance on diets containing phytosterols with 7- and 7,22-configurations, e.g. lathosterol and spinasterol, has only been examined in two species, the generalist H. zea (Ritter and Nes, 1981b) and the specialist P. xylostella (Behmer and Grebenok, 1998) and neither grew particularly well. This may not be unexpected for the specialist, but H. zea feeds on a wide range of plants. Perhaps the preferred host range of H. zea tends to primarily contain 5-sterols, which regardless of side chain configuration, support growth and development (Nes et al., 1997). The ability of lepidopteran larvae to use fungal sterols such as ergosterol is quite variable, even for species within the same family, e.g. the pyralids. This difference seems to reflect general ecology, rather than phylogeny, since caterpillars that can utilize ergosterol tend to be found in environments that favour fungal growth, e.g. damaged fruits and stored products. Highly mobile generalist Lepidoptera larvae are likely to ingest both suitable and unsuitable sterols when they feed. Nes et al. (1997) investigated the consequences of this by rearing H. zea on a synthetic diet that contained
20
S. T. BEHMER AND W. D. NES
different proportions of cholesterol (suitable) and 24-dihydrolanosterol (unsuitable). They found that growth rates began to decrease when the proportion of suitable sterols in the diet dropped below 70%, and that developmental time increased when the proportion dropped below 50%. These results are similar to those found when the grasshopper S. americana was fed diets containing different proportions of usable and non-usable sterols (Behmer and Elias, 1999a, 2000). The only other insects within the Endopterygota to be examined for sterol use are a siphanopteran, Xenopsylla cheopi (Pausch and Fraenkel, 1966) and three hymenopterans. The flea, which feeds primarily on mammals, demonstrated good growth on the phytosterol sitosterol, but did poorly on diets containing stigmasterol and ergosterol. The honeybee, Apis mellifera, in contrast, grew best on diets containing cholesterol and 24-methylenecholesterol, but rather poorly on diets containing other phytosterols (Herbert et al., 1980). The two other Hymenoptera studied were Brachymeria lasus and Pachycerpoideus vindemiae, pupal endoparasites of Lepidoptera and Diptera, respectively (Thompson, 1981). Both larvae grew best on cholesterol, followed by cholestanol, sitosterol and 7-dehydrocholesterol. Cholesterol sparing abilities were also investigated and found to be somewhat limited in both species, but results did suggest that cholestanol worked in this capacity better than did sitosterol or 7-dehydrocholesterol. 3.2
INSECT STEROL METABOLISM
Our understanding of insect sterol use has been greatly enhanced by studies that measure and compare the sterol content of insect tissues to the foods that insects eat. As with sterol use studies there are biases in the existing data, with over half of the species so far examined coming from the Hemiptera and Hymenoptera. Our aim in this section is to describe some of the biosynthetic steps involved in the insect sterol metabolic pathway, and to document how different insect orders metabolize sterols. 3.2.1
Insect sterol metabolic pathways
Insects operate the classic acetate-mevalonate isoprenoid pathway to an unidentified step that forms before synthesis of the 30-carbon olefin, squalene oxide (Fig. 2) (Kircher, 1982; Campbell and Nes, 1983; Silberkang et al., 1983). The post-lanosterol pathway normally operational in other animals is interrupted in insects. For instance, [2-3H]-lanosterol fed to animals is actively converted to cholesterol (Nes et al., 1988b) whereas in the tobacco hornworm, M. sexta, the labelled sterol is absorbed by the gut and metabolized to a sole product, 24,25-dihydrolanosterol, using a reductase-type enzyme (Svoboda et al., 1995a). This enzyme, 24,25-reductase, is also found in the cycloartenolsitosterol pathway of plants (Fig. 3) and has been studied in a cell-free system
INSECT STEROL NUTRITION AND PHYSIOLOGY
21
FIG. 2 Isoprenoid–sterol pathway. HMGR, hydroxymethyl gluturayl CoA-reductase; DXS, 1-deoxy-D-xylulose 5-phosphate synthase; SMT, sterol methyl transferase.
FIG. 3 Generalized plant pathway of cycloartenol conversion to stigmasterol.
of the tobacco hornworm (Short et al., 1996). In plants 24,25-reductase normally recognizes 24-methyl and 24-ethyl desmosterol as substrate to generate campesterol and sitosterol, respectively (Yamada et al., 1997) (Fig. 2), whereas the 24,25-reductase of insects recognizes lanosterol or desmosterol as substrates. The 24,25-reductase enzyme is a critical slow step in the insect conversion of phytosterol to ecdysteroid (Short et al., 1996). As a result, the first inhibitors of sterol metabolism were targeted to inhibit the activity of this enzyme (Svoboda, 1994). Insects, in contrast to more derived animals, have many enzymes in the sterol pathway that can be considered as reverse enzymes that are complementary to
22
S. T. BEHMER AND W. D. NES
those synthesized by plants. For instance, the normal route for conversion of phytosterols involves nucleus changes as follows: the 9,19-cyclopropane group isomerizes to an 8,9-bond, which is then rearranged to the 7-bond. Next a 5bond is introduced and the 7-bond is reduced to form the final 5-structure characteristic of phytosterol end-products (Fig. 3). In contrast, insects can introduce the 7-bond or remove the 5-bond from the sterol nucleus, so that after feeding on plant leaves, the insect will have a sterol composition of 7- or 0-sterols (Svoboda, 1984). For side chain transformations, plants have an enzyme that adds a 22-bond to the sterol side chain, while insects have an enzyme to remove the 22-bond. Likewise, plants convert the 24(25)-bond to form a 24(28)-exomethylene bond while insects convert the 24(28)-bond to a 24,25-double bond. It appears that the catalysis involved with the reverse reactions operated by insects is not simply that of an equilibrium reaction, but rather unique metabolases evolved to carry out the individual reaction. Key enzymes in the post-lanosterol and post-cycloartenol pathway such as a C-4 demethylase, 8,9-isomerase and 23(24)-reductase (in corn) are also absent from insects (Clark and Bloch, 1959b; Ikekawa et al., 1993; Short et al., 1996; Nes et al., 1997). On the basis of present understanding of metabolic relationships, the aliphatic isooctane side chain of cholesterol occupies a position as a key branch point structure of steroid metabolism since it serves as the precursor to ecdysteroids and provides suitable three-dimensional and amphipathic characteristics for sterol to interdigitate in the membrane. Conversion of phytosterol to cholesterol was hypothesized to proceed through a series of side chain metabolisms as outlined in Scheme 2 (Svoboda, 1984) in which the 24-alkyl group is ultimately removed and the resulting 24,25-double bond is saturated to produce the isooctane structure. Sitosterol (7) and stigmasterol (5), the major 24-ethyl cholesterols found in plants, are converted to cholesterol in insects by essentially the same side chain dealkylation pathway. Cell-free systems, originating from both the tobacco hornworm and the silkworm, B. mori, have been used to study C-24 dealkylase (lyase enzyme) and 24,25-reductase (Ikekawa et al., 1993; Short et al., 1996) but unfortunately no detailed characterization or purification of these enzymes has been reported. Therefore, nothing significant is known regarding kinetics, except for reaction mechanisms. Four steps occur in the 24-dealkylation pathway that converts sitosterol to fucosterol, and the first is catalysed by a single dehydrogenase enzyme that lacks stereospecificity. The next step is the stereospecific conversion of fucosterol to the (24R,28R) epoxide of fucosterol. The penultimate step is catalysed by the fucosterol epoxide lyase involving the elimination of acetaldehyde and migration of H-25 to C-24 to form the 24,25-double bond. Finally, in the conversion of desmosterol to cholesterol, a reductase-type enzyme is involved. In related work it was demonstrated that [2-13C]-mevalonic acid is converted to (25S)-[26-13C]-sitosterol (7) (Nes et al., 1992) and, using a cell-free preparation from corn, [27-13C]-lanosterol was converted to
INSECT STEROL NUTRITION AND PHYSIOLOGY
23
SCHEME 2 Pathway of sterol dealkylation by insects and disruption by rationally designed inhibitors targeted for key enzymes in the pathway. The sterol nucleus (SN) in each case is cholesterol based. Structure 7 corresponds to the sitosterol side chain and structure 5 corresponds to the stigmasterol side chain. Structures 10 and 11 correspond to the desmosterol and cholesterol side chains, respectively. Structure 19 is solasodine, a common steroidal alkaloid.
(25R)-[2713C]-24(28)-methylenelanosterol (Nes et al., 1996). This suggested a general process, with the biosynthesis of 24 -ethyl phytosterols passing through three steps: (i) successive methylation at C-24, (ii) isomerization of the 24(28)-bond to the 24,25-bond, and (iii) reduction of the 24,25-double bond – with net retention in configuration at C-25 in the final chiral product. In the case of the biosynthesis of cholesterol via the 24-dealkylation pathway in insects, there is also a net retention in configuration in the final chiral product (Scheme 2). It can be seen that C-2 of mevalonate incorporated into C-26 (a rotation at C-25 is shown to accommodate nomenclature, cf., Guo et al., 1996) is the pro-S-methyl group whereas in cholesterol generated in animals, the pro-R-methyl is derived from C-2 of mevalonate (Popja´k et al., 1977). In the reduction of the 24(25)-double bond, regardless of the system engaged, both hydrogen atoms were added to the si-face of the double bond equivalent to the cis-addition of the hydride ion to C-25 from a pyridine nucleotide and of a proton to C-24. A more precise understanding of the 24-dealkylation pathway was elucidated by the use of structural analogues targeted to disrupt each step. All the inhibitors shown in Scheme 2 block steps along the 24-dealkylation pathway as shown by the accumulation of substrate for the target enzyme. They are also
24
S. T. BEHMER AND W. D. NES
highly effective inhibitors of insect growth (Ikekawa et al., 1993; Short et al., 1996; Svoboda et al., 1994). The mechanism for enzyme inhibition by the ammonium-containing compounds (e.g., 16, 17 and 18) is probably different from the allenes (e.g., 15). It seems that the N-steroids may serve as a highenergy intermediate analogue of the cationic intermediate generated during the catalytic reaction (Short et al., 1996). In contrast, the allenes may serve as a mechanism-based inactivator, much like related compounds prepared and tested with the sterol methyl transferase from yeast (Nes et al., 1998). Natural products, including steroidal alkaloids such as solasodine (19) that contain nitrogen in the side chain, might inhibit the 24,25-reductase as they can inhibit the sterol methyl transferase and inhibit growth of algae by inducing an accumulation of 24-desmethyl sterols with a 24,25-double bond (Mangla and Nes, 2000). However, these N-steroids fail to affect the growth of the tobacco hornworm (Weissenberg et al., 1998), showing important differences between plants and insects in their response to steroidal alkaloids. Prestwich and coworkers (1983, 1984) have studied several side chain modified monofluorinated cholesterols and sitosterols and found that 29-fluorositosterol (20) and 29-fluorostigmasterol were highly toxic to the tobacco hornworm. Abnormal development caused by the 29-fluorosterols is similar to that seen in larvae fed on fluoroacetate. Dealkylation of these fluorosterols produces a lethal product, fluoroacetate, and the inhibitor has no direct effect on phytosterol metabolism. 3.2.2
The Apterygota and Exopterygota
Sterol metabolism has only been examined in four insect orders outside of the Endopterygota. The most primitive is the firebrat, Thermobia domestica, and biochemical studies have shown that it can produce cholesterol from sitosterol (Svoboda and Robbins, 1971). Species in two polyneopteran orders, the Orthoptera and Blattaria, also show the ability to dealkyate. For example, L. migratoria was found to produce radiolabelled cholesterol after it was fed [14C]sitosterol (Rath et al., 1993). Likewise, Costet et al. (1987) and Behmer et al. (1999b) inferred dealkylation of sitosterol in two Schistocerca species reared on diets rich in sitosterol (wheat and artificial diet, respectively) after large quantities of cholesterol were found in the insects’ tissues. However, Behmer et al. (1999b) suggested that S. americana nymphs have a limited ability to dealkylate phytosterols that contain a double bond at the 22-position, e.g. stigmasterol and spinasterol, since these sterols tended to accumulate in the tissues of nymphs reared on diets containing stigmasterol and spinasterol, respectively. Although they also recovered high quantities of cholesterol, it was determined to have originated maternally. Both studies also showed that grasshoppers accumulated 7- and 8-sterols when reared on diets that contained these same sterols, which indicates that grasshoppers also lack the enzymes needed to convert double bonds in the sterol nucleus. Cockroaches,
INSECT STEROL NUTRITION AND PHYSIOLOGY
25
like grasshoppers, produce cholesterol from sitosterol (Robbins et al., 1962), but studies using radiolabelled sterols suggest cockroaches have broader sterol metabolic capabilities than grasshoppers. For instance, Clark and Bloch (1959b) found that the German cockroach produces 22-dehydrocholesterol from ergosterol by saturating the 7–8 double bond and demethylating the side chain. However, they also found that cockroaches, like grasshoppers, could not reduce 22–23 double bonds. Among the Hemiptera, sterol metabolism has been examined in at least one species for each of the three major suborders. The Heteroptera, or true bugs, are the largest of the suborders and have both predaceous and herbivorous members. To date, species representing five superfamilies in this suborder have been investigated for sterol metabolism (Table 3). The two other suborders, Sternorrhyncha and Auchenorrhyncha, contain only phytophagous species and are so far only represented by an aphid and a leafhopper, respectively (Table 3). A comparison of food sterol profiles with insect tissue sterol profiles reveals that in this order, cholesterol is found in aphids (Campbell and Nes, 1983) and a leafhopper (Noda and Koizumi, 2003), but not in any of the phytophagous heteropterans (Table 3). By using [14C]-sitosterol, Campbell and Nes (1983) were able to demonstrate that aphids could dealkylate phytosterols. Noda and Koizumi (2003) have proposed that planthoppers use a 7-reductase and produce 24-methylenecholesterol from ergosta-5,7,24(29)-trienol by intracellular yeastlike symbiotes. Cholesterol is then produced by removal of the side chain group. These results, when combined with the absence of cholesterol in phytophagous heteropterans, suggest that the Heteroptera have lost the ability to dealkylate phytosterols. Meeting sterol requirements, both structural and metabolic ones, has never been considered problematic for predaceous hemipterans since their prey items tend to contain large quantities of cholesterol (Nes and McKean, 1977). In contrast, plant phloem and xylem are not particularly nutrient-rich resources, especially with respect to phytosterols, although it has been demonstrated that phytosterols can be translocated through plants via the phloem (Lehrer et al., 2000; Douglas, this issue). Many phytophagous hemipterans harbour endosymbionts which are known to provide valuable nutrients, especially amino acids (Douglas, 1998), and it has been suggested that these endosymbionts might play a role in sterol nutrition. Initially, it was suggested that they provided sterols directly, but since most bacteria lack the ability to biosynthesize sterols or sterol intermediates this seemed unlikely. Indeed, Campbell and Nes (1983) demonstrated that Schizaphis graminum, an aphid that contains bacterial endosymbionts, did not produce radioactively labelled sterols or sterol metabolic intermediates when it was reared on diets containing [2-14C]-melavonate. Nonetheless, Douglas (1988) has shown that aphid endosymbionts might be linked with cholesterol production. After she reared the green peach aphid, Mzyus persicae, on a holidic diet lacking sterols for five
Dietary sterols (% total)
Insect [diet] Orthoptera (locusts, crickets, grasshoppers and katydids) Acrididae Schistocerca americana (1) [synthetic diet w/ mixed sterols]
Locusta migratoria (2) [wheat] [wheat treated with fungicide]
Heteroptera (true bugs) Reduviidae (I) Rhodnius prolixus (5) [young mice] Arilus cristatus (5) [Tenebrio molitor] Pentatomidae (II) Nezara viridula (5) [mixed diet]a
cholesterol (100.0)
(B) sitosterol (60.0), campesterol (27.0), 22-dihydrobrassicasterol (13.0) (C) stigmasterol (99.0) (D) spinasterol (60.0), 22-dihydrospinasterol (40.0)
cholesterol (81.0), sitosterol (9.4), campesterol (9.4) cholesterol (54.0), stigmasterol (46.0) cholesterol (45.7), spinasterol (37.0), lathosterol (10.3), 22-dihydrospinasterol (6.7) cholesterol (70.0), cholestanol (12.0), sitosterol (6.0), lathosterol (3.5), campesterol (3.5), stigmasterol (1.5) cholesterol (41.5), 8-sterols (24.5), 9 ,19-cyclopropylsterols (21.0), sitosterol (3.0), campesterol (2.0), stigmasterol (1.0), others (7.0)
sitosterol (41.0), stigmasterol (34.0), campesterol (17.0), cholesterol (<1.0)
stigmasterol (34.0), sitosterol (31.0), campesterol (17.0), cholesterol (15.0)
sitosterol, stigmasterol, campesterol
24-methylenecholesterol (56.0), cholesterol (23.0), ergosta-5,7,24(28)-trienol (21.0)
not reported, but mostly cholesterol
cholesterol (98.5), campesterol (1.0)
cholesterol (63.3), 7-dehydrocholesterol (18.4), sitosterol (12.2), campesterol (4.6), stigmasterol (1.5)
cholesterol (92.7), sitosterol (3.5), campesterol (1.4)
sitosterol (48.0–65.7), campesterol (6.0–12.8), stigmasterol (8.0–37.4), isofucosterol (7.7–12.2), C-30 sterols (0.0–12.5)
sitosterol (65.3), campesterol (9.9), stigmasterol (9.8), isofucosterol (8.6), stigmast-7-en-3 -ol (2.8)
S. T. BEHMER AND W. D. NES
Hemiptera (sucking bugs) Sternorrhyncha (psyllids, whiteflies, aphids and coccids) Aphidae Schizaphis graminum (3) [sorghum] Auchenorrhyncha (cicadas, planthoppers and fulgoroids) Delphacidae Laodelphax striatellus (4) [rice]
Insect tissue sterols (% total)
(A) cholesterol (100.0)
sitosterol (54.0), campesterol (24.0), stigmasterol (8.0), isofucosterol (7.0), cycloeucalenol (4.0), obtusifoliol (3.5) 9 ,19-cyclopropylsterols (90.0), 8-sterols (9.0), 5-sterols (1.0)
26
TABLE 3 Sterol profiles from insects and their diets. Insects are organized first by order and then by family. When more than one family occurs within an order, shared roman numerals indicate they come from the same superfamily. No inferences are made about the relatedness of superfamilies to one another.
Acanthosmatidae (II) Cimex lectularius (5) [young mice] Lygaeidae (III) Oncopeltus fasciatus (6) [milkweed] Oncopeltus fasciatus (7) [sunflower] Pyrrhocoridae (IV) Dysdercus fasciatus (8) [rice] Dysdercus cingulatus (6) [cotton seed] Alydidae (V) Megalotomus quinquespinosus (9) [mixed diet]b
Neuroptera (lacewings) Chrysopidae Chrysopa carnea (10) [grain moth eggs] [Schizaphis graminum] Coleoptera (beetles) Dermestidae (I) Trogoderma granarium (11, 12) [synthetic diets]
cholesterol (63.3), 7-dehydrocholesterol (18.4), sitosterol (12.2), campesterol (4.6), stigmasterol (1.5)
cholesterol (88.9), sitosterol (4.1), stigmasterol (4.0), lathosterol (2.4), campesterol (0.6)
not reported, but mostly cholesterol
cholesterol (98.5)
sitosterol (49.1), campesterol (22.3), 24-methylenecholesterol (9.4), isofucosterol (9.1), stigmasterol (8.2) sitosterol (79.5), stigmasterol (10.3), campesterol (10.1)
sitosterol (44.0), campesterol (25.6), 24-methylenecholesterol (11.8), isofucosterol (9.0), stigmasterol (11.8) sitosterol (86.0), campesterol (10.5), stigmasterol (3.2)
sitosterol (97.5), campesterol (2.4), cholesterol (<1.0) sitosterol (91.4), campesterol (4.3), cholesterol (<1.0)
sitosterol (95.1), campesterol (4.8), cholesterol (<1.0) sitosterol (87.6), campesterol (7.0), isofucosterol (3.4), stigmasterol (1.2)
sitosterol (0.0–56.4), spinasterol (0–42.5), 7-stigmastenol (0–39.7), campesterol (0–23.6), stigmasterol (1.2–37.4), isofucosterol (0–9.3), 7-campesterol (0–7.3)
sitosterol (46.6), 7-stigmastenol (13.8), spinasterol (13.4), campesterol (9.0), isofucosterol (5.8), stigmasterol (4.1), 7-campesterol (2.8)
cholesterol (80.2), sitosterol (13.9), campesterol (5.8) sitosterol (54.3), cholesterol (40.3), stigmasterol (5.4)
cholesterol (93.3), campesterol (3.9), sitosterol (2.8) sitosterol (66.8), stigmasterol (19.3), cholesterol (13.8)
(A) sitosterol (69.4), campesterol (25.0), stigmasterol (5.1) (B) [3H]-sitosterol (100.0)
campesterol (59.4), sitosterol (36.7), stigmasterol (2.6), cholesterol (1.3) sitosterol (92.3), campesterol (4.7), cholesterol (<1.0) stigmasterol (85.9), campesterol (5.6), sitosterol (5.4), cholesterol (2.0) desmosterol (93.4), cholesterol (<1.0)
(C) [3H]-stigmasterol (100.0) (D) [3H]-desmosterol (100.0) Coccinellidae (II) Epilachna varivestis (13) [soybean leaves]
sitosterol (55.0), stigmasterol (31.5), campesterol (11.3)
INSECT STEROL NUTRITION AND PHYSIOLOGY
Podisus maculiventris (5) [Tenebrio molitor]
cholestanol (50.7), stigmastanol (20.3), lathosterol (11.8), campestanol (6.0), cholesterol (4.5), other sterols (3.7)
27 (Continued )
28
TABLE 3 Continued Dietary sterols (% total)
Insect [diet] Coccinella septempunctata (14) [aphids] Tenebrionidae (III) Tribolium castaneu (15) [mixed diet]c
Tenebrio molitor (15) [mixed diet]d Curculionidae (IV) Diaprepes abbreviatus (16) [synthetic diet] Hypera postica (17) [alfalfa] Anthonomus grandis (18) [synthetic diet]
not reported
cholesterol (46.4), sitosterol (29.5), campesterol (12.2), stigmasterol (5.5), 7-dehydrocholesterol (3.7), stanols (2.7)
sitosterol (90.2), campesterol (6.5), isofucosterol (2.6)
cholesterol (N – 43.7; I – 4.9), 7-dehydrocholesterol ( þ desmosterol) (N – 39.8; I – 52.3), sitosterol (N – 16.0; I – 25.9), 5,7,24-cholestatrienol (N – 0.0; I – 15.1) cholesterol (66.7), 7-dehydrocholesterol (16.8), sitosterol (11.2), campesterol (3.8)
sitosterol (47.3), isofucosterol (36.7), campesterol (11.8), stigmasterol (4.3) cholesterol (83.0), sitosterol (8.9), campesterol (2.7), 22-dehydrocholesterol (1.6), desmosterol (1.4) spinasterol (69.0), 22-dihydrospinasterol (17.3), avenasterol (13.3) sitosterol (92.0), campesterol (8.0)
(A) [14C]-sitosterol (100.0) 14
(B) [ C]-sitosterol (100.0) þ 25-azacholesterol (C) [14C]-desmosterol (100.0) (D) [14C]-campesterol (100.0) Brachycera (short-horned flies) Muscidae (I) Musca domestica (20) [synthetic diets]
Drosophilidae (II) Drosophila melanogaster (21) [synthetic diets]
cholesterol (90.5), sitosterol (7.1), campesterol (1.9) spinasterol (38.0), lathosterol (34.0), 22-dihydrospinasterol (25.5), avenasterol (11.0) cholesterol (58.1), sitosterol (36.9), campesterol (5.0)
cholesterol (66.1), sitosterol (32.7), campesterol (1.2) sitosterol (53.3), cholesterol (27.7), desmosterol (18.4), campesterol (0.6) cholesterol (97.8), desmosterol (2.0) campesterol (63.4), cholesterol (36.6), sitosterol (trace)
(A) cholesterol (100.0)
cholesterol (99.9)
(B) campesterol (100.0) (C) sitosterol (100.0)
campesterol (95.0), cholesterol (3.3), sitosterol (1.6) sitosterol (90.1), cholesterol (8.1)
(A) cholesterol at 0.1% (100.0)
cholesterol (99.9), others (0.1)
S. T. BEHMER AND W. D. NES
Diptera (flies) Nematocera (long-horned flies) Culicidae Aedes aegypti (19) [synthetic diets]
Insect tissue sterols (% total)
(B) [14C]-desmosterol (100.0) Lepidoptera (moths and butterflies) Pyralidae (I) Plodia interpuntella (23) [mixed diet]e
Galleria mellonella (24) [brood comb] Sphingidae (II) Manduca sexta (25) [synthetic diets]
sitosterol (78.8), isofucosterol (10.4), campesterol (9.3), stigmasterol (1.4)
24-methylenecholesterol (51.9), isofucosterol (21.0), sitosterol (14.2), campesterol (8.0), stigmasterol (4.3) (A) cholesterol (100.0) (B) sitosterol (100.0) (C) stigmasterol (100.0) (D) ergosterol (100.0)
(E) lanosterol (100.0)
(F) obtusifoliol (100.0)
Noctuidae (III) Helicoverpa zea (26, 27) [synthetic diets]
others (97.0), cholesterol (3.0) others (97.3), cholesterol (2.7) others (96.2), cholesterol (3.2) others (95.4), cholesterol (4.6) sitosterol (96.7), campesterol (2.1), cholesterol (1.2) desmosterol (90.3), cholesterol (4.1), sitosterol (2.1), unknowns (3.0)
cholesterol (N – 86.5; I – 14.0), sitosterol (N – 10.8; I – 25.1), desmosterol (N – 1.2; I – 54.2), campesterol (N – 1.2; I – 6.1) cholesterol (85.2), sitosterol (10.7), campesterol (2.5)
cholesterol (87.2), desmosterol (4.4), campesterol (4.3), sitosterol (4.1) cholesterol (77.1), sitosterol (17.4), campesterol (5.5) cholesterol (70.1), stigmasterol (11.2), sitosterol (8.3), campesterol (7.3) cholesterol (49.1), cholest-5,7-dienol (18.2), sitosterol (10.3), campesterol (10.1), ergosterol (7.1), stigmasterol (3.2) dihydrolanosterol (40.1), lanosterol (31.3), cholesterol (18.7), campesterol (3.1), sitosterol (2.9), desmosterol (1.5) obtusifoliol (46.6), cholesterol (35.8), 7-obtusifoliol (6.0), campesterol (3.7), desmosterol (2.8), sitosterol (2.3) cholesterol (100.0)
(B) sitosterol (100.0) (C) stigmasterol (100.0) (D) ergosterol (100.0)
cholesterol (80.0), sitosterol (20.0) cholesterol (84.0), sitosterol (15.0) cholesta-5,7-dienol (41.0), ergosterol (36.0), cholesterol (23.0) clerosterol (80.0), cholesterol (20.0) lathosterol (63.0), spinasterol (34.0), cholesterol (5.0)
29
(A) cholesterol (100.0)
(E) clerosterol (100.0) (F) spinasterol (100.0)
INSECT STEROL NUTRITION AND PHYSIOLOGY
Drosophila melanogaster (22) [synthetic diets]
(B) campesterol at 0.1% (100.0) (C) campesterol at 0.2% (100.0) (D) sitosterol at 0.1% (100.0) (E) sitosterol at 0.2% (100.0) (A) [14C]-sitosterol (100.0)
(Continued )
Continued
Insect [diet]
[synthetic diet w/ mixed sterols]
Noctuidae (III) Helicoverpa zea (28) [corn] [alfalfa]
Spodoptera littoralis (29) [synthetic diet] [synthetic diet with inhibitor]
Tenthredinidae (II) Aneugmenus flavipes (31) [bracken fern]
Dietary sterols (% total)
Insect tissue sterols (% total)
(G) lathosterol (A) cholesterol (90.0), 24-dihydrolanosterol (10.0) (B) cholesterol (70.0), 24-dihydrolanosterol (30.0) (C) cholesterol (50.0), 24-dihydrolanosterol (50.0) (D) cholesterol (30.0), 24-dihydrolanosterol (70.0)
lathosterol (81.0), cholesterol (19.0) cholesterol (93.0), 24-dihydrolanosterol (7.0)
sitosterol (51.0), campesterol þ 22-dihydrobrassicasterol (27.0), isofucosterol (17.0), stigmasterol (6.0) spinasterol (69.0), 22-dihydrospinasterol (17.3), avenasterol (13.3) stigmasterol (100.0)
cholesterol (80.0), campesterol þ 22-dihydrobrassicasterol (11.0), sitosterol (10.0) lathosterol (54.5), 22-dihydrospinasterol (25.5), spinasterol (19.0), avenasterol (0.5)
cholesterol (88.0), 24-dihydrolanosterol (12.0) cholesterol (75.0), 24-dihydrolanosterol (25.0) cholesterol (50.0), 24-dihydrolanosterol (50.0)
stigmasterol (100.0) þ 20,25-diazacholesterol
cholesterol (59.2), desmosterol (15.0), stigmasterol (10.7), sitosterol þ fucosterol (8.5) sitosterol þ fucosterol (25.8), campesterol (22.4), stigmasterol (21.9), cholesterol (11.6), desmosterol (8.0), others (10.3)
sitosterol (83.5), isofucosterol (7.7), 24-methylcholesterol (4.6), cholesterol (3.1)
cholesterol (75.5), desmosterol (13.6), sitosterol (8.0), 24-methylcholesterol (1.5)
sitosterol (93.0), campesterol (7.0)
cholesterol (73.0), sitosterol (17.0), 7-dehydrocholesterol (6.0), campesterol (4.0) cholesterol (lrv – 55.3, adlt – 72.2)*, sitosterol (lrv – 29.5, adlt – 15.7), 7-dehydrocholesterol (lrv – 11.7, adlt – 0.0), desmosterol (lrv – 0.0; adlt – 10.1), 24-methylcholesterol (lrv – 3.5; adlt – 2.0)
sitosterol (87.8), 24-methylcholesterol (8.2), isofucosterol (2.7)
sitosterol (68.7), isofucosterol (16.6), campesterol (9.0), other (3.7), cholesterol (2.0)
cholesterol (73.0), sitosterol (16.6), desmosterol (6.0), campesterol (3.6)
S. T. BEHMER AND W. D. NES
Hymenoptera (sawflies, wasps, bees and ants) Symphyta (sawflies) Xyelidae (I) Macroxyela ferruginea (30) [elm] Diprionidae (II) Neodiprion pratti (31) [pine] Neodiprion lecontei (30) [pine]
30
TABLE 3
Argidae (II) Atomocera decepta (30) [Hibiscus] Cephidae (III) Cephus cinctus (30) [wheat] Xiphydriidea (IV) Xiphydria maculata (31) [fungus]
Braconidae (I) Microplitis demolitor (32) [H. zea reared on cholesterol diet] [H. zea reared on ergosterol diet] Apidae (II) Apis mellifera (33) [maple]
Apis mellifera (33) [goldenrod]
Megachilidae (II) Megachile rotundata (34) [alfalfa] Anthophoridae (II) Diadasia rinconis (35) [three Sonoran cacti – average values]
INSECT STEROL NUTRITION AND PHYSIOLOGY
Dolerus nitens (31) [fescue grass]
sitosterol (62.0), campesterol (21.3), isofucosterol (4.4), stigmasterol (3.5), other (7.7), cholesterol (1.1)
cholesterol (56.6), sitosterol (17.6), desmosterol (13.6), campesterol (7.6), lathosterol (2.4), other (2.2)
sitosterol (55.3), stigmasterol (31.0), 24-methylcholesterol (10.8), 7-dehydrocholesterol (2.9)
cholesterol (68.5), sitosterol (10.8), trans-22-dehydrocholesterol (10.1), 24-methylcholesterol (6.8)
sitosterol (50.2), 24-methylcholesterol (22.4), stigmasterol (11.4), isofucosterol (10.1), cholesterol (5.9)
cholesterol (72.7), 24-methylcholesterol (14.6), sitosterol (10.3), 7-dehydrocholesterol (1.6)
stigmasterol (32.9), sitosterol (26.8), stigmastanol (12.9), campesterol (7.9), 22-stigmastenol (7.8), 7-stigmastenol (5.8), campesterol (1.9), cholesterol (1.2), 7,22-stigmastadienol (1.1)
7-dehydrocholesterol (74.9), cholestanol (7.9), other (8.8), cholesterol (5.7), ergosterol (2.3)
cholesterol (100.0)
cholesterol (100.0)
7-dehydrocholesterol (56.0), ergosterol (27.0), cholesterol (14.0), 22-dihydroergosterol (3.0)
7-dehydrocholesterol (76.0), cholesterol (24.0)
isofucosterol (55.1), sitosterol (18.3), 24-methylenecholesterol (12.7)
24-methylenecholesterol (pp – 55.8; adlt – 41.7)*, isofucosterol (pp – 19.0; adlt – 22.0), sitosterol (pp – 13.2; adlt – 12.7), cholesterol (pp – 0.6; adlt – 0.7) 24-methylenecholesterol (pp – 52.6; adlt – 34.6)*, 7-stigmasten-3 -ol (pp – 13.2; adlt – 20.4), isofucosterol (pp – 12.6; adlt – 21.8), sitosterol (pp – 9.8; adlt – 12.4), cholesterol (pp – 1.0; adlt – 2.0)
7-stigmasten-3 -ol (66.2), sitosterol (5.7), isofucosterol (5.6), 24-methylenecholesterol (1.5)
not reported
Isofucosterol (40.7), 24-methylenecholesterol (34.1), sitosterol (13.0), campesterol (10.4)
24-methylenecholesterol (84.2), sitosterol (4.3), cycloartenol (3.1), fucosterol (2.8), campesterol (0.5)
24-methylenecholesterol (92.2), sitosterol (1.3), cycloartenol (0.0), fucosterol (2.4), campesterol (1.7)
31
(Continued )
32
TABLE 3 Continued Dietary sterols (% total)
Insect [diet] Vespidae (III) Dolichovespula maculata (34) [omnivorous] Vespula maculifrons (34) [omnivorous] Formicidae (III) Atta cephalotes (36) [fungal garden]f Acromyrmex octospinosus (37) [fungal garden]g
Solenopsis invicta (34) [culture diet]h
Formica exsectoides (34) [not reported]
not reported
not reported
not reported 5,7-4-desmethylsterol (gngyl – 84.6; myclm – 75.4), 5-4-desmethylsterol (gngyl – 1.5; myclm – 3.0), 7-4-desmethylsterol (gngyl – 8.4; myclm – 10.9), unknown sterols (ngyl – 5.5; myclm – 10.7) not reported
not reported
Insect tissue sterols (% total)
cholesterol (86.5), campesterol (3.8), sitosterol (3.7), desmosterol (3.1), 24-methylenecholesterol (0.6) cholesterol (81.5), sitosterol (7.8), desmosterol (5.8), campesterol (2.6), 24-methylenecholesterol (0.6) 7-dehydro-24-methylenecholesterol (70.0), 22-dihydroergosterol (20.0), ergosterol (10.0) 5,7-4-desmethylsterol (pp – 94.1; wrkr – 84.7) þ, 5-4-desmethylsterol (pp – 1.2; wrkr – 8.3), 7-4-desmethylsterol (pp – 1.8; wrkr – 3.2), unknown sterols (pp – 2.9; wrkr – 4.0) cholesterol (pp – 69.5; adlt – 45.4),** campesterol (pp – 23.9; adlt – 26.0), sitosterol (pp – 3.8; adlt – 17.4), isofucosterol (pp – 2.3; adlt – 5.2) cholesterol (69.4), desmosterol (9.4), campesterol (5.5), sitosterol (3.9)
S. T. BEHMER AND W. D. NES
(1) Behmer et al., 1999b; (2) Costet et al., 1987; (3) Campbell and Nes, 1983; (4) Noda and Koizumi, 2003; (5) Svoboda et al., 1984; (6) Svoboda et al., 1984; (7) Svoboda et al., 1976; (8) Gibson et al., 1983; (9) Feldlaufer et al., 1986; (10) Keiser and Yazlovetski, 1988; (11) Svoboda et al., 1980a; (12) Svoboda et al., 1980b; (13) Svoboda and Thompson, 1973; (14) Svoboda and Robbins, 1978; (15) Svoboda and Lusby, 1994; (16) Ward et al., 1991; (17) MacDonald et al., 1990; (18) Earle et al., 1967; (19) Svoboda et al., 1982; (20) Feldlaufer and Svoboda, 1991; (21) Feldlaufer et al., 1995; (22) Svoboda et al., 1989a; (23) Svoboda and Lusby, 1994; (24) Feldlaufer et al., 1997; (25) Svoboda et al., 1995a; (26) Nes et al., 1997; (27) Ritter, 1984; (28) MacDonald et al., 1990; (29) Svoboda et al., 1989b; (30) Svoboda et al., 1995b; (31) Schiff and Feldlaufer, 1996; (32) Ritter and Johnson, 1991; (33) Svoboda et al., 1983; (34) Svoboda and Lusby, 1986; (35) Feldlaufer et al., 1993; (36) Ritter et al., 1982; (37) Maurer et al., 1992. a Sunflower seeds, peanuts and green beans. Reported values indicate the range of specific sterols in the available foods. b Soybean, sunflower, and alfalfa seeds, with fresh green beans and water. Reported values indicate the range of specific sterols in the available foods. c White flour/whole wheat flour/extracted brewers yeast (44:44:12) þ 0.1% sitosterol. N ¼ normal diet, I ¼ diet with sterol inhibitor (500 ppm 25-axacoprostane). d Commercial bran meal (no added sterol). e White corn meal/whole wheat flour/glycerol/honey/extracted brewer’s yeast/rolled oats/wheat germ (28:26:17:16:6:4:3), plus 0.05% sitosterol; N ¼ normal diet, I ¼ diet with sterol inhibitor (2 ppm 25-axacoprostane). f From oak leaves and oatmeal bran (values reported for the gongylidia (gngly) and mycelium (myclm)). g From brambles and privet plants. h A mixture of housefly prepupae, boiled eggs and honey/water (1:1). * lrv=larvae; adlt=adult. ** pp=pupae; adlt=adult. þ pp=pupae; wrkr=worker.
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generations, she found that the reproductive effort of these aphids was similar to those that had been reared on an identical diet supplemented with cholesterol. However, when she disrupted the endosymbionts using the antibiotic chlortetracycline, her ‘‘aposymbiotic’’ aphids only produced viable offspring when they were reared on the holidic diet that contained cholesterol. She suggested that the bacteria in the aphids might have a sparing effect on the insect’s requirement for sterols, although she did not speculate on how such a mechanism might be operating. 3.2.3
The Endopterygota
Sterol metabolism has been examined in 27 endopterygotes, representing predaceous and phytophagous lifestyles across five different orders. In the one neuropteran studied, the common green lacewing, Chrysopa carnae, insect tissue sterol profiles closely mirrored those found in the food (Keiser and Yazlovetski, 1988). This indicates that little sterol metabolism was occurring, which is not surprising considering that most larval neuropterans are predaceous. Among the Coleoptera that have been examined, which includes phytophagous and predaceous species from the suborder Polyphaga but no representative species from the smaller suborders Archostemata, Myxophaga or Adephaga, no clear pattern of sterol metabolism emerges and in some cases the findings have been quite surprising. For example, the kharpa beetle, Trogoderma granarium, seems to lack the ability to dealkylate phytosterols even though it is a stored-products pest (Svoboda et al., 1980a,b). Coccinella septempuncta also seems to lack the ability to dealkylate sterols (Svoboda and Robbins, 1978), but this is somewhat expected since most coccinelids are predaceous. However, the Mexican bean beetle, Epilachna varivestis, is unique among the coccinelids in that has become secondarily phytophagous and is able, to some degree, to dealkylate sterols (Svoboda and Thompson, 1973). Additionally, this species is unusual because it reduces the 5–6 double bond in the sterol nucleus and has a tissue sterol profile dominated by saturated sterols (Svoboda and Thompson, 1973). The remaining beetle species that have been studied are all phytophagous, and while they all show an ability to dealkylate, other aspects of their sterol metabolic capabilities vary. For instance, both the tenebrionids studied can convert 5-phytosterols to cholesterol, but they also tend to produce and accumulate 7-dehydrocholesterol (Svoboda and Lusby, 1994). Among the curculionids, tissue sterol profiles vary depending on the type of dietary sterols present. For example, the boll weevil, Anthonomus grandis, feeds on cotton and is capable of producing cholesterol from sitosterol, although it also tends to accumulate a large quantity of unmetabolized sitosterol (Earle et al., 1967). In contrast, the alfalfa weevil, Hypera postica, does not produce cholesterol and appears to have a limited capacity to dealkylate the C-24 side chain (MacDonald et al., 1990). Alfalfa plants only
34
S. T. BEHMER AND W. D. NES
contain phytosterols with 7,22-configurations, which suggests that a double bond at the 22-position may prevent dealkylation in this insect, and that metabolism of the sterol nucleus is not possible. Whether this is the case for most phytophagous beetles is unclear. Flies from three different superfamilies have now been analysed for sterol metabolism, and the results suggest that dealkylation only occurs in the Nematocera (long-horned flies), which are considered primitive to the Brachycera (short-horned flies). For instance, when larvae of the yellow fever mosquitoes, A. aegypti (Nematocera), were reared on an artificial diet containing [14C]-sitosterol, [14C]-desmosterol or [14C]-campesterol, radiolabelled cholesterol was always recovered (Svoboda et al., 1982). In contrast, when the brachyceran M. domestica was fed radiolabelled [3H]-sitosterol, no radiolabelled cholesterol was identified from the insect tissue (Kaplanis et al., 1963, 1965). Radiolabelled sterols were also used to demonstrate the lack of dealkylation in a second brachyceran, the fruit fly D. melanogaster (Svoboda et al., 1989a). Even Drosophila species that are specialists on cacti lack the ability to dealkylate phytosterols, as demonstrated by Kircher et al. (1984). They analysed the sterol profiles of D. mojavensis and D. nigrospiracula and found that they closely matched the phytosterol profile of their respective hostplants, and contained only minor amounts of cholesterol. Much of the pioneering work into sterol metabolism in insects was conducted on two lepidopteran species, M. sexta (reviewed by Svoboda and Weirich, 1995) and B. mori (reviewed by Ikekawa et al., 1993), and we have reported some of the key details from these studies in the preceding section. In general, lepidopteran insects are capable of producing cholesterol from C28 and C29 5-phytosterols (Table 3). In some species, however, a double bond at the 22-position and/or a methyl group rather than an ethyl group at C-24 seems to reduce the efficiency of this conversion (e.g. Svoboda and Robbins, 1968; Svoboda et al., 1995a). Metabolism of the sterol nucleus has only been studied in two species, M. sexta (Svoboda et al., 1995a) and H. zea (Ritter, 1984, 1986; Nes et al., 1997), and results suggest it does not occur. When these two species were reared on diets containing 0-, 5-, 7- and 8-sterols, the sterol nucleus double bond configuration found in the caterpillars resembled that found in the diet. Cholesterol was also found in all of these caterpillars, but it was probably maternal in origin. Finally, the Hymenoptera seem to show a sterol metabolic pattern similar to the Diptera. That is, dealkylation is present in the primitive members of the order but is lost in the more derived members. For instance, Schaefer et al. (1965) demonstrated that the Virginia sawfly, Neodiprion pratti, could convert [3H]-sitosterol to [3H]-cholesterol. Likewise, cholesterol is the dominate tissue sterol found in most Symphyta even though the plants on which they feed contain primarily 5-phytosterols (Table 3). The one exception to this is Xiphydria maculata, which is thought to feed on fungus and has tissue sterol profiles dominated by 7-dehydrocholesterol (Svoboda et al., 1995b). Among
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35
the predaceous Apocrita, which primarily includes the wasps and some ants, cholesterol is found in the tissues in large quantities, along with various other phytosterols that are normally associated with the dietary sterols of their prey items (Table 3). This also seems to be the case for the hymenopteran parasitoid, Microplitis demolitor (Braconidae) (Ritter and Johnson, 1991). Interestingly these authors found that the sterol composition of this parasitoid’s host had little effect on its growth. However, growth was effected when they reduced the amount of cholesterol available in the haemolymph of the host. This suggests parasitoids, especially those found in herbivorous insects, have effective cholesterol sparing mechanisms above a minimal cholesterol threshold. Among the ants that tend fungal gardens, the finding that their sterol composition usually mirrors that of the fungus indicates that little sterol metabolism occurs (Maurer et al., 1992). This is an interesting finding because it perhaps suggests that the central nervous system can function in certain organisms without cholesterol. Finally, among the bees that have been examined, 24-methylenecholesterol is the dominant tissue sterol, followed by other phytosterols that are determined by the sterol composition of the bee’s host-plants. Regardless of the dietary sterol available to worker bees, 24-methylenecholesterol is the major tissue sterol found in the brood reared by workers (Svoboda et al., 1983). This is accomplished through a selective transfer of sterols from the endogenous sterol pools of the workers to the developing larvae through the brood food material secreted from the hypopharyngeal and mandibular glands and/or the honey stomach of the workers (Svoboda et al., 1986). 3.3
CONSIDERATIONS OF PATTERNS OF STEROL USE AND METABOLISM
Taken together information about sterol use (Table 2) and metabolism (Table 3) can give useful insights into the evolution of sterol abilities in insects, especially when viewed in a phylogenetic context (Fig. 4). Cholesterol supports growth for most insects, but since it is not always readily available, many insects, especially plant feeders, must dealkylate phytosterols to produce cholesterol. This conversion also provides the necessary precursor to ecdysone, which, along with 20-OH ecdysone, is one of the two most commonly occurring moulting hormones (ecdysteroids) in insects (Svoboda, 1994). That a thysanuran can convert sitosterol to cholesterol suggests the ability to dealkylate the phytosterol side chain had an early evolutionary origin (Fig. 4a). This ability is maintained in some of the Polyneoptera, as demonstrated by the fact that grasshoppers and cockroaches convert sitosterol to cholesterol. The Orthoptera are the best studied exopterygotes, and it is interesting to note that within this group the grasshoppers (acridids), as compared to the crickets (gryllids), have a very limited ability to dealkylate. The Califera (the suborder to which the acridids belong) are derived from the Ensifera
36
S. T. BEHMER AND W. D. NES
FIG. 4 A Phylogenetic interpretation of sterol metabolic capabilities in the major insect orders. The phylogeny was adapted from the Tree of Life (Maddison et al., 2001), and represents Kristensen’s (1991) cautious views about the relationships of orders. Dealkylation is considered the primitive state based on the ability of a thysanuran to convert sitosterol to cholesterol (see Section 3.2.2, p. 24). Bars across phylogenetic branches indicate a lost ability to dealkylate. Question marks have been placed next to bars where we hypothesize that dealkylation abilities might be lost, but have yet to be tested. The triple question mark for the Strepsiptera indicates the placement of this branch within the Endopterygota remains uncertain.
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(the suborder containing the gryllids) (Rowell and Flook, 1998), which suggests that broad sterol metabolic dealkylation abilities may have been lost. It also perhaps implies that there may be metabolic costs associated with maintaining a broad enzymatic capability. Unfortunately there is little consensus on the phylogenetic relationship within the Polyneoptera (Exopterygota excluding the Apterygota and Paleoptera), and the lack of information on sterol use or metabolism in this group, outside of the acridids, makes it difficult to infer much about the evolution of sterol metabolic capability. Based on our current understanding of the relatedness and feeding habits of the Polyneoptera, we suggest that the loss of dealkylation has occurred independently on multiple occasions. Since the polytomy for this group is soft, clarification of the relationships between the major orders within the Polyneoptera will likely indicate fewer independent losses of sterol dealkylating ability. The hemipteroid assemblage includes the Hemiptera, Thysanoptera, Psocoptera and Phthiraptera, but in this clade only hemipteran species have been examined. The Hemiptera are, however, a large group and inspection of the sterol metabolism data for this group reveals an interesting pattern. The primitive Sternorrhyncha and Auchenorrhyncha can dealkylate phytosterols, but the more derived Heteroptera cannot. Predaceous feeding is the primitive state in the Heteroptera, while plant feeding has arisen secondarily. This result suggests that once dealkylating abilities are lost they may be difficult to regain. An additional unique aspect associated with the loss of dealkylation ability in this group is the use of different moulting hormones by the primitive predaceous hemipterans (C27 ecdysteroids) and their more derived plant-feeding cousins (C28 (Makisterone A) or C29 (Makisterone C) ecdysteroids). Where predaceous feeding has arisen secondarily among the plant feeders, however, Makisterone A is retained as the moulting hormone (Aldrich et al., 1982; Svoboda et al., 1984). This suggests that the mechanisms for ecdysteroid biosynthesis are strongly conserved among the more derived Heteroptera even when cholesterol is abundant (Svoboda, 1994). Based on the feeding lifestyle of the Phthiraptera, we propose that dealkylation abilities may have been lost in this group. Phytophagy appears to be the primitive condition in Coleoptera and it seems that most phytophagous beetles have a broad ability to dealkylate, except perhaps for those, such as T. granarium, which have adopted a plant-feeding lifestyle secondarily. The coccinellid E. varivestis, which has also evolved plant feeding secondarily, is a slight exception. It can dealkylate sitosterol but not other phytosterols, including campesterol. The Neuroptera, Megaloptera and Rhaphidiodea are found in the same clade as the Coleoptera (Fig. 4b), but there is evidence that the ability to dealkylate may have been lost in these members, which would not be surprising considering their predaceous nature. The basal members of the Hymenoptera, Lepidoptera and Diptera are phytophagous, and dealkylation is found in all of the primitive members of
38
S. T. BEHMER AND W. D. NES
these orders. So far, it appears that this ability has been maintained throughout the Lepidoptera, but not throughout the Hymenoptera or Diptera. Among the Hymenoptera, dealkylation ability has been lost in the Apocrita, so bees, like the derived hemipterans, use various dietary sterols in unmetabolized form and have Makisterone A as their moulting hormone. Likewise, fungus-eating ants do not metabolize their dietary sterols, e.g. ergosterol, and contain primarily C28 ecdysteroids (Maurer et al., 1991, 1993). Finally, the derived Diptera (Brachycerca) also appear to have lost the ability to dealkylate, and biochemical studies with M. domestica and D. melanogaster revealed that these two derived Diptera produce Makisterone A from campesterol (Feldlaufer and Svoboda, 1991). Nonetheless, these same authors found that C27 ecdysteroids were the major moulting hormones and suggested that they were probably derived from trace amounts of cholesterol in the diet, which were most likely obtained through selective uptake. The predaceous nature of the Mecoptera and Siphonaptera also lead us to suggest that dealkylation abilities may be lost in these two groups, but whether this has occurred independently in each group depends on resolving the relationship of these two Orders to one another.
4
Insect sterol physiology
Why cholesterol accumulates in some but not all insects is puzzling. Moreover, it is unclear why insects have evolved enzymes that transform sitosterol to cholesterol, but not ones that transform cycloartenol and other intermediates of the phytosterol pathway to cholesterol. To examine these enigmas the function of sterols in insects needed to be uncovered. At present sterols are considered to play three roles in insect physiology: (i) as a membrane insert; (ii) as a precursor to the moulting hormones, ecdysteroids; and (iii) as a signalling molecule bound to the hedgehog group of proteins affecting development. Interestingly, the sterol features and amounts involved with each of these functions differ. The bulk of the sterol in insects is localized in microsomes, a combination of membranous systems (Short et al., 1996), which is in agreement with the proposal that the major role for sterols is as an architectural component of membranes (Clark and Bloch, 1959a; Nes and McKean, 1977). Although some insect species have rigid sterol structural requirements, others appear to be able to use a variety of different sterols under different environmental conditions (Behmer and Elias, 1999a, 2000; Mondy and Corio-Costet, 2000). However, when the level of required sterols falls below a critical threshold, for example from 350 g/ml to 55 g/ml in L. migratoria nymphs that have been reared on wheat with a modified sterol profile, insect growth and embryogenesis are impaired (Costet et al., 1987). In an analogous finding, humans that possess inborn errors of cholesterol metabolism accumulate sterol intermediates. This
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39
accumulation corresponds with a drastic reduction in plasma cholesterol levels – normal, 160 mg/dl, diseased, less than 110 mg/dl – and children that express the error in cholesterol metabolism often exhibit physical malformations (Tint et al., 1997). The total amount of sterol found in an insect is influenced by its size, and generally the larger the insect the more sterol it contains. For example, aphids possess ca. 0.02 g/insect while the tobacco hornworm contains ca. 1800 g/larva (Table 4). The sterol composition of an insect depends on its diet and for phytophagous insects a mixture of cholesterol and 24-alkyl sterols is common (95% of total sterol). The size of the insect in relation to the amount of sterol in the diet is also important. Thus, to grow and complete development the corn earworm requires ca. 200 ppm of added sterol (Nes et al., 1997) whereas the tobacco hornworm requires ca. 320 ppm of added sterol (Svoboda et al., 1995a). In these two studies the total amount of sterol delivered to individual larva was approximately 1–3 mg, which is about the amount of sterol (ca. 1 mg) in a mature sorghum leaf (Nes, 1990). The natural distribution of sterols isolated from insects suggest that they must possess three key features to serve as a membrane insert, including a 3 -hydroxyl group, a flat tetracyclic ring system and an 8–10 carbon side chain positioned in the ‘‘right-handed’’ conformation. The structure of phytosterols juxtaposed as shown in Fig. 5 fit the dimension of the monolayer of the lipid leaflet (Parker and Nes, 1992), but based on the type of sterols accumulated in insects, it seems that some cannot tolerate significant variations in the shape of the cholesterol molecule. Tolerance to variation in sterol structure has been examined in numerous insects by replacing cholesterol with other sterols that either lack a particular feature, e.g., 4,4-dimethyl group, or have some other interesting aspect to them e.g., altered stereochemistry or contained an ‘‘extra’’ methyl group in the sterol side chain (Clark and Bloch, 1959a; Clayton, 1964; Ritter and Nes, 1981a,b; Kircher, 1982; Nes et al., 1982, 1997; Ikekawa et al., 1993; Behmer and Elias, 1999a, 2000). In summary, these structure–activity studies revealed that intermediates in the sitosterol–cholesterol pathways that had methyl groups at C-4, or possessed a 9,19-cyclopropane group or 8-bond could not be utilized by insects. The insect membranes can use sitosterol together with cholesterol, since the methyl group found on sitosterol does not lengthen the side chain beyond functional limits. In addition, the natural stereochemistry at C-20 in phytosterols gives necessary directionality to the overall length of the molecule, thereby contributing to its planar shape, as shown in the conformational perspective illustrated in Fig. 5. In support of this proposal, when cholesterol and either 20-epicholesterol or 17(20)Z-dehydrocholesterol were fed to the silkworm, only cholesterol supported growth (Ikekawa et al., 1993). The stereochemistry of the 24-alkyl group in phytosterols is of less importance to membrane structure, since both sets of compounds are utilized equally well by the insect and undergo metabolism to ecdysteroids.
40
TABLE 4 Total sterol amount for species from different orders. Numbers in parentheses indicate the cited study. Species
Growth form Ametabolous – no metamorphosis Thysanura – primitively wingless, e.g. firebrat Hemimetabolous – incomplete metamorphosis Orthoptera – straight winged, e.g. grasshoppers Homoptera – winged and wingless, e.g. aphids Hemiptera – half-winged, e.g stink bugs Holometabolous – complete metamorphosis Hymenoptera – membrane winged, e.g. ants Coleoptera – forewings thickened, e.g. beetles Lepidoptera – scale-covered wings, e.g. moths
*
NE* NE Schistocerca americana (1) Schizaphis graminum (2) NE Solenopsis invicata (3) Anthonumus grandis (4) Helicoverpa zea (5) Manduca sexta (6) NE
20.00 0.02 2.00 55.00 66.00 1800.00
(1) Behmer et al., 1999b; (2) Campbell and Nes, 1983; (3) Ba et al., 1995; (4) Nes, unpublished; (5) Nes et al., 1997; (6) Svoboda et al., 1995a. NE, not examined.
S. T. BEHMER AND W. D. NES
Diptera – one pair of wings, e.g. flies
Total sterol (g)
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FIG. 5 Conformational perspective of cholesterol.
The importance of metabolizing the 24-alkyl(alkylidene) group to the cholesterol side chain can be found in the structure of 20-hydroxecdysone (Fig. 6). The moulting hormone requires a 25-hydroxyl group for activity and the 24-alkyl group in the phytosterol side chain will, due to steric factors, interfere with the introduction of the 25-hydroxyl group. The importance of converting phytosterol intermediates to a 5-4-desmethyl sterol is also related to the structure and function of ecdysteroids (Gilbert et al., 2002). The removal of geminal groups at C-3 of cycloartenol is required for a sterol to possess an unhindered 3-hydroxyl group. The elimination of the C-4 methyl groups is necessary for sterol–lipid interactions (Nes et al., 1978) and to covalently bind to protein during Hedgehog signalling (Porter et al., 1996). An insect responds, presumably through the Hedgehog signalling process, to subtle differences in the structure of the side chain, which suggests a greater level of sterol recognition by the insect-signalling mechanisms than is currently recognized in other animal systems. Thus, corn earworm cultured on a synthetic diet treated with non-utilizable sterols, such as ergosta-5,23-dienol, proceed through a very long pupation stage and the adults which emerge are abnormal with malformed wings and legs (Nes et al., 1997). Future efforts to establish the substrate features and quantitative importance of sterol for specific functions during growth and maturation of individual insect species will prove to be a formidable task. However, the use of inhibitors and genetic engineering approaches to assist in providing sufficient protein or enzymes that act on steroids may provide answers. The aim of the remainder of this section is to document the journey of dietary sterols through an insect. We start at the mouth and describe what is known about the ability of insects to taste sterols and regulate their intake via behavioral mechanisms. We next turn our attention to the fate of sterols after they are ingested. The digestive process begins in the mouth and continues as the food is passed through the foregut into the midgut. During this process
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FIG. 6 Roles for cholesterol. Diagrammatic representation of cholesterol functions – (A) membrane insert, (B) precursor to the moulting hormone 20-hydroxyecdysone, and (C) as a component of the hedgehog signalling process. See text for details.
sterols can either be modified, via enzymatic reactions, or remain unchanged. Because we have already discussed many of the details related to sterol metabolism, we focus on the processes involved in their uptake and absorption. We now know that transport of sterols through the haemolymph is facilitated by lipophorin, and we touch on this process before turning our attention to the tissue and intracellular distribution of sterols in an insect. We also examine the central role sterols play in various physiological processes, including development, growth, and reproduction, as well as their influence on parasites and viruses of insects. 4.1
STEROL TASTE AND THE REGULATION OF INTAKE
Whether insects can directly taste sterols has only critically been examined in grasshoppers. In two studies using S. americana nymphs, Champagne and Bernays (1991) and Behmer et al. (1999a) used a series of short-term experiments to explore how different sterols affected feeding behavior, such as
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initial contacts with different test foods and the length and structure of the first meal. Results from these experiments suggested that this grasshopper could not taste sterols, even after they had fed for 24 h on spinach, a plant that is devoid of suitable sterols. Perhaps this finding is not surprising considering the strong lipophilic nature of sterols and the design and manner in which contact chemoreceptors operate. However, until detailed electrophysiological work has been performed, the question of whether insects can directly taste sterols will remain unanswered. Despite the lack of evidence indicating an ability to directly taste sterols, S. americana nymphs still seem capable of adjusting their feeding behavior in response to the sterol profile of their food. For example, when grasshoppers were given spinach, the lengths of successive meals declined until the food was completely rejected (Champagne and Bernays, 1991). This aversion response was eliminated, however, when sitosterol or cholesterol was applied topically to the spinach leaves. Notably the length of the first meal on spinach was identical whether or not a suitable sterol had been added, which indicated there was no immediate effect of sterol profile on feeding behavior. These overall results suggest that grasshoppers were behaving in a fashion that is consistent with food aversion learning mediated through post-ingestive feedbacks (e.g. Lee and Bernays, 1988). That sterols might be responsible for this learned aversion was investigated in a series of experiments by Behmer and Elias (1999b). First, they compared feeding behavior of grasshoppers on artificial diets that contained sitosterol or a spinach lipid extract, and found that the diet containing the extract evoked a deterrent feeding response. Next, they separated the spinach lipid extract into three classes: desmethyl sterols, dimethyl sterols and remaining spinach lipids. These were then added to artificial diets and feeding behavior was again analysed. This experiment showed that only the desmethyl sterol extract, which contains the spinach end-product sterols, deterred feeding. In a separate set of experiments, Behmer et al. (1999a) tested whether the deterrent response to sterols was mediated post-ingestively. In one experiment grasshoppers were given an initial meal on spinach and then injected, through the abdomen, with a small volume of mineral oil that contained ‘soybean’ sitosterol (suitable), ‘spinach’ sterols (unsuitable) or no sterol (the control). Following the injection the grasshoppers were returned to their arenas and given a fresh leaf of spinach and the length of their next meal was recorded. Results showed no difference in feeding time between the ‘soybean’ sitosterol and no sterol treatment (about 75 s each), but showed a significant drop in the meal length of the ‘spinach’ sterol injected grasshoppers (median feed time of only 3 s). It was suggested that the grasshoppers were developing a learned association between the presence of an unsuitable sterol in the blood with the taste of the spinach. This was tested, and verified, by giving grasshoppers artificial foods containing different combinations of sterols and flavors.
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The feeding responses of beetles and caterpillars to sterols have also been examined, although in less detail, and the results are mixed. Among the beetles, for example, cholesterol and sitosterol stimulate biting, but not feeding, in Leptinotarsa decemlineata (Hsiao and Fraenkel, 1968), but neither biting nor feeding in Sericesthis geminata (Wensler and Dudzinski, 1972). Shanks and Doss (1987) examined five weevil species, and found that sitosterol and stigmasterol stimulated feeding in only one of these, the obscure root weevil, Sciphithis obscurus. In the same study a synergistic effect of sterols and sucrose on feeding was observed in S. obscurus as well as in the black vine weevil, Otiorhynchus sulcatus, and the pea leaf weevil, Sitona lineatus. The silkworm, B. mori, is the most thoroughly studied caterpillar with regards to the role of sterols in feeding behavior. Hamamura et al. (1962) reported that sitosterol stimulated feeding, but Nayar and Fraenkel (1962), using commercially obtained preparations of sitosterol of fair purity, found no such effect. Likewise, they reported no effect on feeding by melissyl alcohol, cholesterol, sitosterol, or stigmasterol. However, Ito et al. (1964) showed that silkworm feeding could be stimulated if commercial sitosterol was recrystallized. The functional significance of this is difficult to interpret, however, since they also showed that recrystallizing stigmasterol and campesterol produced a similar effect. Finally, Hamamura (1970) studied biting responses of B. mori towards sterols and measured high levels of activity towards sitosterol but less towards cholesterol and only a few towards ergosterol. Chippendale and Reddy (1972b) showed that in Diatraea grandiosella, cholesterol and sitosterol failed to stimulate biting or feeding. They did show, however, that feeding was negatively effected by three cholesterol esters (three different acid moieties (acetate, myristate, oleate)), which suggests that a substituted 3 hydroxyl group might adversely affect the insect’s sensory mechanisms. We suggest, however, that the interpretations from the beetle and caterpillar be viewed cautiously since the assays used did not look at detailed aspects of individual behavior. 4.2
STEROL ABSORPTION
The midgut appears to be the main site of sterol absorption for many insects, particularly in phytophagous species (Joshi and Agarwal, 1977), as was recently demonstrated by Jouni et al. (2002a). In their study they placed a block of food containing [3H]-cholesterol in the foregut, midgut or hindgut of 5th stadium M. sexta larvae and, after 2 h, measured the radioactivity of the haemolymph. Results showed that when the food was placed in the midgut, radioactivity levels in the haemolymph were 5-fold and 9.5-fold higher compared to when the food was placed in the foregut and hindgut, respectively. In some phytophagous insects, specialized regions of the midgut may be particularly active in sterol absorption. For example, 6 h after Joshi and
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Agarwal (1977) fed cabbage leaves containing [3H]-cholesterol to the grasshoppers, S. gregaria and Hieroglyphus nigrorepleteus, they found that the gastric caeca contained more radioactivity (>50%) than any other region of the gut. In contrast, the same study indicated that in omnivorous species such as the house cricket, Gryllodes sigilatus (Orthoptera: Gryllidae), the carpenter ant, Camponotus compressus (Hymenoptera: Formicidae) and a diving beetle, Dytiscus (Coleoptera: Dytisidae), the foregut, especially the crop, was the primary site of sterol absorption. A similar finding was reported for the cockroaches (Blattaria: Blattidae) Euryocotis floridana (Clayton et al., 1964) and Periplaneta americana (Joshi and Agarwal, 1976), which are also omnivorous. The mechanisms governing the uptake of sterols from the food to gut tissues in insects are poorly understood. In vertebrates, the absorption of sterols into intestinal cells is aided by bile salts. Insects do not have bile salts and current working models suggest that sterols, packaged as mixed micelles with phospholipids, are absorbed passively into the membranes of gut tissue (Turunen and Crailsheim, 1996). In contrast to insects, vertebrates show a strong tendency to absorb cholesterol more easily than phytosterols. In humans for example, cholesterol absorption is 56.2 12.1% in normal subjects during test meals (Ostlund, 2002), but the values for phytosterols are much lower, ranging from 0.04% for sitostanol (Ostlund et al., 2002) up to 16% for campesterol (Lutjohann et al., 1995). This later study also suggests that humans absorb 5-phytosterols more easily than they do 0-sterols (stanols). Sterol uptake mechanisms seem less rigid in insects however, as evidenced by the fact that phytosterols are regularly found in the tissues of both predaceous and phytophagous insects (Table 3). There are some exceptions though, as Robbins (1963) has demonstrated with houseflies, which can selectively absorb and incorporate cholesterol, relative to other dietary sterols, when it is limiting in the diet. Phytosterols reduce cholesterol uptake in humans and for this reason phytosterols are considered beneficial dietary components even though they are non-usable. Phytosterols also reduce cholesterol uptake in the zoophagous D. maculatus, but this negatively effects growth since this species requires a dietary source of cholesterol (Katz et al., 1971). Behmer et al. (1999b) recently explored how usable and non-usable sterols interact to influence sterol absorption and cholesterol production in phytophagous insects. They fed S. americana nymphs artificial diets containing ‘usable’ sitosterol (0.5 g/mg), ‘non-usable’ stigmasterol (0.5 g/mg) or a mixture of these two sterols (equal amounts for a total of 1.0 g/mg) and then compared the tissue sterol profiles. The results showed that the relative amount of stigmasterol in the tissue of ‘mixture’ grasshoppers was significantly lower compared with the ‘stigmasterol’ grasshoppers, even though both treatments contained similar total amounts of stigmasterol. The results also showed that the relative cholesterol level from ‘mixture’ grasshoppers was intermediate between the ‘usable’ sitosterol and
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‘unusable’ stigmasterol treatments. This suggests that phytophagous insects probably do not selectively absorb usable over non-usable sterols, but that non-usable sterols can affect cholesterol production, most likely by reducing the uptake of sterols that serve as precursors to it. In general, it seems that the amounts of sterol absorbed, as well as the amount of cholesterol produced, are directly related to the proportions of usable and non-usable sterols in the food. Other aspects of an insect’s food also influence sterol absorption. For example, Dupnik and Kamm (1970) showed that dietary lipids enhanced the efficiency of sterol uptake in the grass moth Crambus tricestus (Lepidoptera: Crambidae), and Ito and Nakasone (1966) demonstrated that soybean fatty acids increased the utilization of sitosterol in B. mori. Plant secondary metabolites, on the other hand, may reduce sterol utilization, as demonstrated by Weissenberg et al. (1998). They fed M. sexta larvae diets containing the alkaloid spirosolane and glycoalkaloids and found that sterol absorption and metabolism was significantly reduced. However, the negative effect of these alkaloids decreased as the concentration of cholesterol and sitosterol in the food increased, as shown by Weissenberg et al. (1998) with the beetle T. castaneum and Bloem et al. (1989) with the caterpillars H. zea and S. exigua. Sterols can enter and leave midgut mucosal cells rather quickly, as demonstrated by Jouni et al. (2002a). They fed a block of food containing [3H]-cholesterol to 5th instar M. sexta larvae and found that radioactivity levels in the midgut peaked within the first hour of feeding. This peak was then followed by a sharp decrease within 4 h, after which there was a steady decrease until late pupation, at which point the tissue started to disintegrate. Numerous studies have shown that insects can absorb dietary cholesterol both in the free and ester form, but that following absorption a small fraction of the free form is subjected to intracellular esterification (Turunen and Chippendale, 1977; Kuthiala and Ritter, 1988; Komnick and Giesa, 1994; Jouni et al., 2002a). Esterification takes place in intestinal microsomes, fat body and ovaries via acyl coenzyme A: cholesterol acyltransferase (ACAT), but the rate at which exogenous sterols are esterified is affected by various side chain modifications and the number and position of double bonds in the B-ring of the sterol nucleus (Billheimer et al., 1983; Macauley et al., 1986). The significance of esterification is unknown, but it has been suggested that it could serve as a temporary intracellular storage mechanism (Turunen, 1985). Few studies have examined the rates of uptake of radiolabelled phytosterols. Phytophagous insects, compared to zoophagous or omnivorous ones, rarely encounter cholesterol in their diets and in fact most of the sterols they encounter are present as fatty acid esters. We advocate the use of radiolabelled phytosterols and phytosterol esters as a method for more properly investigating sterol absorption in phytophagous insects. It would also lead
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to a more thorough understanding of the manner in which different phytosterols interact with each other, plus other diet constituents, in the insect midgut. 4.3
STEROL TRANSPORT AND TISSUE DISTRIBUTION
Following absorption into the midgut, sterols must be transported through the haemolymph to organs of utilization and storage. However, the hydrophobic nature of sterols requires that they be carried through the cytosol and haemolymph by a specialized carrier system. The specific mechanisms by which cholesterol and other sterols are delivered from the haemoceol through the midgut to different tissues and organs are poorly understood. Recently, however, a putative sterol carrier protein thought to be involved in intracellular transport of cholesterol has been identified in the yellow fever mosquito, A. aegypti (Krebs and Lan, 2003). Likewise, high-density lipophorin (HDLp), a non-covalent mixture of lipid and protein, has been identified as the sole haemolymph carrier of cholesterol in M. sexta (Jouni et al., 2002a). 4.3.1
Intracellular transport
The identification of a putative sterol carrier protein involved in the intracellular transport of cholesterol is significant because it provides a mechanism by which sterols can be delivered to the endoplasmic reticulum (ER), the organelle where sterol dealkylation and ecdysteroid production are thought to occur. Krebs and Lan (2003) identified the putative mosquito sterol carrier protein-2 (SCP-2) using cDNA from a fourth instar subtracted cDNA library of A. aegypti and the sterol transfer domain AeSCP-2 was shown to have a high degree of homology in the sterol transfer domain to both rat and human SCP-2 (69%). Transcripts of AeSCP-2 differed between 4th instar larvae and early pupae, being strongly detected in larval midgut tissues, but not in the head and hindgut. This outcome is consistent with the midgut being the primary centre of intracellular cholesterol trafficking in immature insects. In contrast, AeSCP-2 transcripts in the pupae were mainly observed in the thorax, head and body wall of the abdomen, but not in the gut. When interactions between AeSCP-2 and cholesterol were examined, using a radiolabelled cholesterol-binding assay, purified recombinant AeSCP-2 showed a high affinity to cholesterol. They also found that the C-terminal of AeSCP-2 did not have the peroxisome targeting sequence (AKL or SKL) found in vertebrate SCP-2 members. This led them to speculate that AeSCP-2 may be involved in the transfer of sterol between cellular compartments such as lysosomes, ER, plasma membrane and mitochondria. Interestingly they showed that the addition of 60 ng/ml of 20-OH ecdysone to an A. aegypti tissue culture (primarily gut tissue) increased AeSCP-2 expression, which suggests that ecdysteroids may be directly involved in regulating cholesterol uptake in
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the midgut of insects. The only other SCP-2 family member so far described in insects is the Drosophila SCP-X (X97685), but surprisingly the amino acid similarities between AeSCP-2 and SCP-2 domain in Drosophila are low (52%) (Krebs and Lan, 2003). 4.3.2
Haemolymph transport
The discovery that HDLp is the sole carrier of sterols through the haemolymph (Jouni et al., 2002a) is also important because without it cholesterol could not be transferred from sites of absorption or synthesis to sites of storage or utilization. Lipophorin was first identified in the 60s and is best known for its role in the transport of diacylglycerols (DG) between midgut tissues, the fat body and flight muscles, where DG is ultimately used to fuel flight (Ryan and van der Horst, 2000). HDLp is one of three existing forms of lipophorin, each being distinguished from one another based on their densities (Beenakkers et al., 1985). High-density lipophorin (HDLp), which carries sterols, contains relatively little lipid, while low-density lipophorin (LDLp) contains more. A third class, the very-high-density lipophorin (VHDLp) is sometimes produced, and its most common form is vitellogenin. Lipophorin, which tends to be spherical in shape, has a surface composed of phospholipids and proteins and an internal core that is mainly diacyglycerol (DAG) plus smaller amounts of sterols, hydrocarbons, carotenoids and other acylglycerols (Soulages and Wells, 1994). It is produced in the fat body but resides in the haemolymph, where it acts as a reusable shuttle that can easily interface with various tissues to collect and transport cholesterol throughout the body (Chino, 1985). The mechanism by which DG and cholesterol are transported from the midgut to target tissues appears to differ fundamentally. Whereas DG transport is mediated by the HDLp receptor and lipid transfer particle (LTP) in both the midgut (Canavoso and Wells, 2001) and fat body (Canavoso et al., 2003), cholesterol transfer between the midgut and fat body through HDLp seems to operate via aqueous diffusion (Yun et al., 2002). Such a mode of delivery for cholesterol is supported by two findings. First, the transfer of cholesterol from the midgut to HDLp, and from HDLp to fat body shows saturation kinetics. Second, treating the midgut and fat body tissues with suramin, a known inhibitor of fat body HDLp receptor (Tsuchida and Wells, 1990), enocytic inhibitors, anit-LTP antibodies or EDTA, has little or no effect on cholesterol transfer. Based on these results, Yun et al. (2002) proposed that in M. sexta the maintenance of cholesterol homeostasis takes place by a mass action mechanism where cholesterol is freely transferred between lipophorin and tissue depending on the needs of the tissues. Additionally they found that the transfer of cholesterol between fat body and HDLp was bi-directional, but that the transfer between midgut and HDLp was uni-directional (midgut to HDLp). They suggest this might be related to relatively high cholesterol content
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of the midgut basolateral membrane, which would favour the export of cholesterol, but inhibit its import. The fat body is effectively a storage centre for a number of molecules, especially lipids, and in a separate study Jouni et al. (2002b) explored cholesterol efflux from the fat body to HDLp in late instar M. sexta larvae. They demonstrated that lipophorin specifically and significantly induced a transfer of [3H]-cholesterol from the fat body, and then conducted a number of experiments to determine whether cholesterol transfer from fat body to HDLp was receptor-mediated or followed an aqueous diffusion pathway. Several results, including a high Km, high activation energy and the lack of Ca2 þ dependence favoured the aqueous diffusion model. They also examined whether LTP or lipophorin receptor was involved in cholesterol transfer, and found that anti-LTP inhibited cholesterol transfer by 20%, while sumarin, which inhibits lipophorin receptor, reduced cholesterol transfer by 50%. These last two results indicate that LTP-dependent and receptor-mediated pathways may be involved in the transfer process of cholesterol from fat body to HDLp, but Jouni et al. (2002b) argue that if these pathways exist, they likely represent minor routes. Experiments using cholesterol-labelled lipophorin have also been conducted to investigate the fate of lipophorin-cholesterol. Jouni et al. (2002a), using 5th instar M. sexta larvae, found that injected cholesterol was cleared from the haemolymph with a half-life of 10.2 h. After 17 h they found that 30% of the radioactivity remained in the haemolymph but that 38% of it was recovered in the fat body. Only 11% was recovered from the midgut, which agrees with the finding that transfer of cholesterol between the midgut and HDLp is uni-directional towards HDLp (Yun et al., 2002). 4.3.3
Tissue and intracellular distribution
Cholesterol is the dominant tissue sterol for most insects but few detailed studies on the distribution of cholesterol in various insect tissues exist. Lasser et al. (1966) found that total sterol concentration of the tissues from the skunk roach, E. floridana, were similar to, but slightly lower than the concentrations of sterols found in the tissues of vertebrates. Upon inspecting the sterol concentration of individual organs from adults they found that cholesterol concentrations were highest in nerve tissues and salivary glands, intermediate in the gut, Malpighian tubules and fat body, and lowest in muscle and cuticle. Interestingly, increasing the dietary concentration of cholesterol 10-fold, to 1.0%, resulted in a 7-fold increase in the cholesterol content of the fat body. This strongly suggests that the fat body can act as a storage site for excess cholesterol. Other tissues showed significant, but much smaller increases. Tissue sterol profiles can also differ between males and females, and across developmental stages. For instance, Lasser et al. (1966) showed that there was
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a 3-fold higher concentration of cholesterol in the reproductive organs of females compared with males. With respect to development, Ba et al. (1995) detected 26 sterols in the imported red fire ant collected from field stations in Texas and found that an individual ant’s sterol composition was influenced by the stage of development – egg, larva, worker or queen. Eggs contained the least diversity in type of sterol, with cholesterol dominating the total sterol mixture (45% total sterol), whereas the queen possessed the greatest sterol diversity with 24-alkyl sterols detected and 24-ethyl sterols, such as sitosterol, dominating the sterol mixture (54% total sterol). Larvae contained many of the sterols found in the egg, with cholesterol being the most abundant (50% total sterol). Minor amounts of unusual sterols were also isolated, including ergosta-5,23-dienol (3 in Scheme 1) which is produced as a natural phytosterol in corn (Guo et al., 1995). There is strong evidence that tissue, especially nerve tissue, exhibits selective uptake of cholesterol when it is limiting. For example, when Lasser et al. (1966) reared E. floridana on a diet of 0.005% cholesterol and 0.1% cholestanol, they found that brain tissue had a 1:1 ratio of cholesterol:cholestanol, but that the ratio of these two sterols in the other tissues, such as the cuticle (1:4) and the alimentary canal (1:10), were much lower. Likewise, Dwivedy (1993) found that 6-times more cholesterol than sitosterol was recovered from brain tissues of M. domestica larvae reared on a diet containing cholesterol (0.002%) and sitosterol (0.01%). In contrast, all other tissues, e.g. gut, fat body and muscle, from these houseflies had greater proportions of sitosterol than cholesterol. This suggests that nerve tissues might have special quantitative cholesterol requirements compared to other tissues and that the idea of a cholesterol sparing mechanism may extend beyond metabolic, i.e. ecdysteroid, functions to also include specific tissue requirements. It would be interesting to see whether similar selective uptake of cholesterol occur in phytophagous insects with limited dealkylating capabilities when they are reared on foods containing mixtures of sterols that can and cannot be dealkylated. Recently it has been shown that in mammalian membranes, cholesterol is not uniformly miscible, but rather has a tendency to organize the bilayer into cholesterol-rich liquid or cholesterol-poor liquid disorded domains. This leads to the formation of specialized phase domains called rafts, which are known to play an important part in both polarized protein sorting and signal transduction (Simons and Ikonen, 1997; Brown and London, 1997). Rietveld et al. (1999) explored whether these rafts also exist in D. melanogaster, and found that although Drosophila membranes had ergosterol as the predominant membrane sterol (69%), they still contained rafts with similar protein and lipid composition. They also identified a detergent-insoluble fraction in the Drosophila membranes that, like mammalian rafts, was rich in sterol, sphingolipids and glycosylphosphatidlyinostil-linked proteins, and demonstrated that the sterol-linked Hedgehog N-terminal fragment associated specifically with this detergent-insoluble membrane fragment. This, they
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suggest, demonstrates that raft formation is preserved across widely separated phyla in organisms that contain different lipid structures. Lipid rafts have also been found in two caterpillars, H. virescens and M. sexta, but in contrast to Drosophila, cholesterol was the dominant sterol (Zhuang et al., 2002). The proportion of esterified sterols found in different insect tissues can also vary depending on the sterol composition of the diet, as well as the insect’s developmental stage and sex. Lasser et al. (1966) found that the nymphal tissues of E. floridana reared on a low-cholesterol/high-cholestanol diet contained cholesterol primarily in the free form, and that a large proportion of the cholestanol from these same tissues occurred in the ester form. This same study also found that adults had a much higher percentage of esterified sterols in their tissues, even when they were reared on a diet containing only cholesterol. Jouni et al. (2002a) reported a similar finding with M. sexta, but noted a more dramatic shift from the free form to the esterified form in males. In their study male larvae and pupa contained only minor quantities of esterified cholesterol, but in adults the majority of sterols recovered from the fat body were esterified. Female larvae and pupae, in contrast to males, contain sizeable pools of sterol esters, particularly in the fat body. At the adult stage, however, female fat body sterol profiles are quite similar to males. Such results might reflect the fact that in adults the demand for unesterified cholesterol to occupy specific functional spaces is reduced compared to during growth, when a limited availability of cholesterol in the diet might lead to a strict confinement of cholesterol to this space. There is also evidence for sterol turnover in insects (Vroman et al., 1964; Lasser et al., 1966), but the rate at which it occurs depends on the concentrations and types of sterols in the food. In general, the half-life of free cholesterol is significantly longer than that of the free sparing sterol pool, and the greatest turnover of sterol occurs in the alimentary canal. To our knowledge, the subcellular distribution of sterols has only been examined in one insect, the skunk roach E. floridana (Lasser and Clayton, 1966). In this study roaches were reared on a ‘cholesterol-sparing’ diet (0.005% cholesterol þ 0.1% cholestanol) and, after reaching maturity, various samples of muscle, salivary glands and nerve tissue were removed and separated into ‘nuclear’, ‘mitochondrial’, and ‘microsomal’ fractions by differential centrifugation. Sterol concentrations were found to be highest in the microsomal and mitochondria fractions and lowest in the nuclei fraction, but in all three fractions the pool of free sterols was larger than the pool of sterol esters. Three emergent sterol peaks corresponding to cholesterol, cholestanol and 7-cholestanol (a metabolite of cholestanol) were identified and results showed that no single sterol was confined to any particular subcellular fraction. Lastly, they showed that the proportions of these three sterols in each of the fractions within a tissue type were similar, but across the different tissues the proportions varied.
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4.4
S. T. BEHMER AND W. D. NES STEROL REPRODUCTIVE PHYSIOLOGY
A common observation for many insects is that regardless of the sterol in the diet, cholesterol is the dominate sterol that accumulates in eggs (Dutky et al., 1963; Ichimasa, 1976; Lubzens et al., 1981; Costet et al., 1987; Ba et al., 1995; Behmer et al., 1999b; Jouni et al., 2002a). This suggests that cholesterol might be functionally significant. The three-dimensional structure (length– width–volume) of cholesterol (Fig. 5) shows that it has flatness (a series of parallel planes containing all the atoms), as determined by the transfusions of the ring junctions and the existence of the equatorial attachments of a polar head (3 -hydroxyl group), and a non-polar tail (17 -side chain). Phytosterols share many of the same three-dimensional characteristics, as indicated in the X-ray structures shown in Fig. 7. However, the presence of a bulky 24-alkyl group is assumed to increase the fluidity of the bilayer as a result of altered phase-transitions which causes the side chain to sweep out a modified cone structure compared to the cone structure swept out by the cholesterol side chain (Fig. 7). The significance of cholesterol in eggs may also relate to the fact that it serves as a precursor to numerous ecdysteroids, which regulate a number of critical processes throughout embryonic development (Sall et al., 1983). Almost half of the sterol found in eggs is esterified (Dutky et al., 1963; Ichimasa, 1976; Jouni et al., 2002a), and Ichimasa (1976) found that in the silkworm, the accumulation of sterol esters was higher in ‘diapause’ than in ‘non-diapause’ ovaries. The mechanisms governing the transfer of cholesterol to eggs from the fat body via lipophorin, and the contribution of LTP to this process have recently been investigated using the silkworm, B. mori (Jouni et al., 2003). In this study pupal ovarioles (5-day) were incubated with [3H]-cholesterol-labelled
FIG. 7 X-ray structures showing biosynthetic interrelationships of cycloartenol, sitosterol and cholesterol.
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lipophorin under a number of different conditions. Results showed that cholesterol transfer exhibited hyperbolic dependency on lipophorin concentration, with a Km of 0.83 0.17 mg/ml, and that cholesterol transfer was strongly temperature dependent. The role of LTP in cholesterol transfer from lipophorin to ovarioles was studied by treating the ovarioles with anti-LTP IgG and then incubating them with cholesterol-labelled lipophorin. This had no effect on the transfer of cholesterol, indicating that LTP was not involved. Taken together, these findings suggest that cholesterol transfer between lipophorin and ovarioles follows an aqueous diffusion mode of action, which is similar to other studies that have investigated the process of cholesterol transfer between tissues in insects (Jouni et al., 2002a; Yun et al., 2002). The particular importance of cholesterol to developing embryos was demonstrated elegantly by Costet et al. (1987) when they reared newly moulted adult female L. migratoria on wheat seedlings that contained 9 ,19cyclopropylsterols in place of normal 5-sterols (following treatment with fungicide). This caused a shift in the tissue sterol profile of the females, from mostly 5-sterols (81%), to a mixture of 5-sterols (47%), 8-sterols (24.5%) and 9 ,19-cyclopropylsterols (21%). Females on the modified wheat also saw a 90% reduction in cholesterol titres of their haemolymph compared with controls. These differences had no effect on the normal egg-laying cycles, a phenomenon also observed in flies reared on cholesterol-free diets (Monroe, 1959), but they did find that egg production in the locusts reared on the modified wheat was reduced by 10–20% compared with the control group. Not surprisingly, they found that egg sterol profiles reflected the dietary sterol profile, with control eggs containing 95.5% 5- and 0-sterols plus a small fraction of 7-sterols, and experimental eggs containing a mixture of 38% cholesterol, 35.5% 8-sterols, and 24.5% 9 ,19-cyclopropylsterols. Ecdysteroid content of the eggs from the two treatments also varied, with controls, on average, containing 3-fold higher concentrations. They also discovered that the ecdysteroid values of eggs within a pod were similar, so for each pod they destructively sampled 3–4 eggs and then measured percentage hatch for each pod. Their results indicated that when ecdysteroid content fell below 70–80% of the controls, no development occurred, perhaps because no meiotic reinitiation took place. When titres were 50–70% below normal, embryonic development was initiated, but the serosal cuticle was only partially laid down (or not at all) and little normal swelling of the egg occurred. At 40–50% below normal, some eggs hatched normally but many of them had embyros with complex malformations. If, however, ecdysteroid titres were no more than 40% below normal, embryogenesis was normal and viable larvae hatched. Unfortunately, this study cannot completely disentangle the effects of cholesterol from ecdysteroids. Nonetheless, the abnormalities found in the developing embryos would suggest that the lack of cholesterol in adult females, which was needed to produce ecdysteroids for the eggs, was
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probably a greater factor in the low observed viability than was the low cholesterol titre in the eggs. Finally, the effects of dietary sterol, and maternal sterol allocation, on reproduction were examined in the diamondback moth, P. xylostella (Behmer and Grebenok, 1998). In this study caterpillars were reared on diets containing one of three phytosterols, sitosterol, stigmasterol or spinasterol, and two control diets (a ‘wheatgerm’ control, and a trace sterol control). When the females eclosed they were collected and mated with males that came from a culture reared on the ‘wheatgerm’ diet. Fecundity and egg viability were measured and then five second-day neonates from 10 different females were reared on the maternal diet. These were reared and mated with culture males, and once again fecundity and egg viability were measured. Results demonstrated that dietary sterols affected egg viability but not total egg production. This seems to be a recurring phenomenon in insects, and suggests that the mechanisms driving egg production might be independent from those determining egg viability, which is clearly linked to dietary sterols, either directly, or indirectly via ecdysteroid production. There was also a suggestion that the lack of large differences in viability between some of the treatments may have been the result of sterol contributions by males to females during copulation. The males used for mating were reared on diets containing sitosterol, which is readily converted to cholesterol by the diamondback moth. Nuptial gifts have been reported in a number of lepidopteran species (Dussourd et al., 1991; Wiklund et al., 1993; Sculley and Boggs, 1996) but whether sterols are part of this contribution has not been reported.
5
Insect sterol ecology
It seems highly unlikely that sterol requirements would influence the food selection behavior of predaceous insects since cholesterol is probably never a limiting nutrient. Likewise, specialist insect herbivores, by definition, should be specifically adapted to the sterol profiles, concentration and spatial distribution of sterols found in their host-plants, e. g. some of the cactophilic Drosophila species (Fogleman et al., 1986). On the other hand, it has been suggested that constraints on sterol metabolic capacity may be a contributing factor driving food mixing behavior in generalist insect herbivores (Bernays, 1992; Behmer and Elias, 2000). Diverse plant communities provide a rich tapestry of dietary sterols, but little is known about the functional significance of variation in plant sterol profiles. One possibility is that sterol profiles may reflect adaptations to local abiotic conditions. For example, plants in the Chenopodiaceae, which are characterized by high concentrations of 7-desmethylsterols (Xu et al., 1988), are found in alkaline prairies of the United States of America, saline habitats around the Mediterranean, Caspian and Red seas and as weeds in salt-rich soils around human habitations
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(Heywood, 1993). Likewise, 7-desmethylsterols are the major sterols found in the Cucurbitaceae, another family that has members specifically adapted to the dry conditions of the desert (Heywood, 1993). It is, however, also possible that phytosterol profile may function as a unique defence against insect herbivores. Grasshoppers are often found in abundance in dry, saline habitats, and it is interesting to note that chenopods and cucurbs often dominate where large grasshopper communities occur, yet these plants do not suffer great damage (Chambers et al., 1996, personal observations). That phytosterols might function as a defence is strengthened by the finding that grasshoppers have very sensitive behavioral responses, including learning, to food sterol profiles (Champagne and Bernays, 1991; Behmer and Elias, 1999b; Behmer et al., 1999a). Traditionally, the terms specialist and generalist refer to the host range of an insect herbivore, but it could also be used to describe an insect’s sterol utilization abilities. However, host and sterol use range is not always correlated. For instance, D. pachea is a specialist in both senses because it feeds only on senita cactus rots and can only grow on very specific sterols. In contrast, the hornworm, M. sexta, is a specialist on plants in the Solanaceae, but it is a sterol generalist since it is capable of growing on a wide range of sterols (Svoboda and Robbins, 1968). That M. sexta has such a broad metabolic capability despite its restricted diet is probably explained by the fact that solanaceous plants contain a suite of different sterols. In complete contrast to M. sexta are the grasshoppers, which are primarily generalist feeders but sterol specialists since sitosterol is the only phytosterol they can convert to cholesterol (Behmer et al., 1999b). This does not mean that they must have a diet consisting only of sitosterol, but results from experiments where foods contained mixtures of sterols indicate that their capacity to spare cholesterol is much reduced compared to cockroaches (Lasser and Clayton, 1966) and hide beetles (Clark and Bloch, 1959a). Unfortunately the early findings in cockroaches and hide beetles, plus work on flies slightly later (Cooke and Sang, 1970), led most researchers to believe that sterol sparing mechanisms were probably common in insects. While very few studies have explicitly explored this ability in insects, the work on grasshoppers, plus a study using H. zea (Nes et al., 1997), suggest that such an ability may be highly constrained in generalist mandibulate insect herbivores. Perhaps a need to maintain high cholesterol levels partially explains cannibalistic behavior that is observed in some phytophagous insects. There is good evidence in humans that plant sterols can inhibit the process of carcinogenesis and reduce the risk of developing certain cancers (Rao and Korathkar, 1997). Likewise, studies with insects have shown that sterol nutrition can influence an insect’s ability to deal with plant defences and viruses. For example, Bloem et al. (1989) showed that the addition of tomatine to the diet of H. zea significantly reduced growth and food utilization in a linear dose-dependent fashion. However, when they added equimolar
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cholesterol to the same diet, growth and food utilization levels in H. zea returned to normal. Phytophagous insects are often observed eating plant reproductive tissues, such as petals and pollen, that are rich in cholesterol, and it would be interesting to test whether an insect, as a way of self-medicating, might select a diet rich in cholesterol after ingesting tomatine. Dietary sterols can also affect attacking viruses, as demonstrated by MacDonald and Ritter (1988). They found that the sterols in a single-nucleocapsid nuclear polyhedrosis virus (HzSNPV) were similar to those of H. zea, the host insect, and that the HzSNPV isolated from the larvae fed 5-, 0-, or 5,7-sterols contained primarily cholesterol, cholestanol or 7-dehydrocholesterol, respectively. Theirs was the first study to report an insect or animal virus containing sterols other than cholesterol or desmosterol. They also found that dietary sterol influenced virulence, and that the LD50 of viruses originating from insects reared on a lathosterol diet was lower compared with those collected from insects reared on diets containing cholesterol, cholestanol or ergosterol. Ritter (1988) also examined the effects of dietary sterols on an entomogenous nematode. She reared Steinernema feltiae on a diet supplemented with various sterols and found that development was good on 5-, 7- and 0-sterols, but suffered on diets containing 5,7- or 22-sterols. However, when cholesterol was added to the 5,7-sterol diet, growth recovered to normal. Perhaps in some insects, maintaining a specific sterol tissue profile might provide a novel form of resistance against nematodes. Little is known about how dietary sterol of an insect host influences ectoparasitic mites, but the neutral sterol profile of Varroa jacobsoni was found to match that of its honey bee host, Apis mellifera (Feldlaufer and Hartfelder, 1997). Surprisingly, V. jacobsoni is unable to convert dietary sterols to cholesterol, yet it still produces significant amounts of C27 ecdysteroids (Feldlaufer and Hartfelder, 1997).
6
Applied implications and evolution of sterol metabolic constraints
Billions of dollars are spent throughout the world each year trying to reduce the impact of insect herbivores on food crops, yet insect herbivores continue to cause severe problems and at the same time are capable of rapidly evolving resistance to commonly used pesticides. Designing control methods around insect sterol metabolic constraints may, however, provide a unique and powerful approach that specifically targets insects, while leaving non-target organisms, especially vertebrates, unaffected. Recent advances in biochemical and genetic approaches now make it possible to manipulate the sterol compositions of plants to examine how such modifications affect insect growth and development. For example, Costet et al. (1987) used a fungicide to modify the sterol composition of wheat (Costet et al., 1987) while others
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(Corbin et al., 2001; Nes et al., unpublished data) have engineered plants by introducing foreign genes of the sterol pathway. This can generate a modified sterol composition of non-utilizable sterols, such as increased levels of 4-methyl intermediates or side chain olefins such as ergosta-5,23-dienol. The traditional approach of searching for novel dealkylation inhibitors should also lead to the discovery of highly specific insecticides that are non-toxic to plants as well as humans, e.g. targeting an inhibitor for the enzyme catalysing the 24-dealkylation pathway, which would be insect-specific. We feel that by recognizing the natural differences in sterol ecology among insect herbivores, it may be possible to design plants or prepare inhibitors for use against individual species, per se, rather than for broad-spectrum use. The evolution of insect resistance to various insecticides is documented throughout the literature, and it is interesting to speculate about whether insects could evolve rapidly in response to the presence of novel phytosterols that have been bioengineered into their host-plants. At present, no study exists exploring the evolution of sterol metabolic capacities in any insect herbivore, but Behmer and Grebenok (1998) have data that suggest genetic variation for sterol use and metabolism within populations of insect herbivores does exist. Working with P. xylostella, they found lower survival and reduced performance for larvae reared on stigmasterol diets compared with those reared on sitosterol-based diets. Additionally, within the stigmasterol population, variation existed among individuals in larval and pupal development time, mass gain and fecundity. This was a key finding because it indicated that the genetic variation existed upon which selective pressure could act. Somewhat surprisingly, evidence was found suggesting that selection may already be occurring in the second-generation stigmasterol-reared caterpillars. For example, larval and development time was reduced compared with the first generation caterpillars, and there was an increase in both the number of viable eggs produced and in the overall percent viability in the second generation. It would have been interesting with measure various performance measurements over successive generations on the stigmasterol diet to see whether the responses would, after sufficient time, be similar to control reared insects. Additionally, it would be interesting to see, through the application of quantitative genetics methods, whether there were any costs associated with having evolved novel sterol use and metabolic capabilities.
7
Conclusions
It remains a mystery why insects, as well as other arthropods, have lost the ability to synthesize cholesterol, especially considering its fundamental role in many biological processes. Without doubt we have come a long way in our understanding of sterol requirements and metabolism in insects, but a more systematic approach is needed if we are to fully comprehend the relationship
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between insects, particularly phytophagous ones, and their dietary sterols. In this regard we advocate the use of a phylogenetic framework as the guiding principle for selecting groups of insects for future studies on sterol metabolism, and where possible, sterol use. Our hope is that this will lead to the discovery of the type and amount of sterols that are species-specific and host-specific, and that it will shed light on their origins and evolution. In particular we advocate targeting insect groups that show transitions from phytophagy to predaceous lifestyles, followed by phytophagy arising secondarily. This is because such groups provide independent contrasts to study the loss and/or gains of dealkylating ability. We are also starting to gain a greater appreciation for how insects use sterols once they are ingested, and the mechanisms which are employed to regulate their intake, metabolism, transport and distribution within and between different tissues and organelles. However, a stronger foundation in our understanding of the metabolic pathway by which insects generate cholesterol is needed if we are to understand how cholesterol homeostasis in insects is achieved. The role of sterols in cellular signalling processes should also be characterized, and questions of regulation of enzyme activities and gene expression in response to development, and to environment, need to be examined, preferably by way of a comparative approach. Studies exploring how insect sterol physiology interfaces with ecology are in their infancy, but there is evidence to suggest that such interactions could be playing a role in modifying behavior, especially food selection in phytophagous insects. For example, how important are phytosterols in determining patterns of feeding among generalist phytophagous insects? To what extent must generalist insects with sterol metabolic constraints limit their intake of unsuitable phytosterols, and what amounts are tolerable? Are sterols limiting for phytophagous insects, either during development or for reproduction? Perhaps a greater understanding of the relationship between sterols and insects will shed new light on how insect–plant interactions are shaped. Insect sterol metabolic constraints also represent a unique opportunity for bioengineering, and offer a highly target-specific approach to pest management. One of the immediate challenges is to elaborate the specific details, enzymologically and otherwise, of the metabolism of sterols and to identify sites in the pathway, or in absorption/transport, that can be targeted for rational drug design. It is also our hope that generating transgenic plants with modified sterol compositions, such as alterations in the intermediate to endproduct ratios, or specific modifications in the 24-alkyl sterol side chain structures, could be used to study plant–insect interactions, including the evolution of sterol metabolic capabilities. The study of insect sterol nutrition and physiology is a compelling enterprise, and we think that future studies on sterol use in insects will continue to reveal new and fascinating discoveries, with important implications at both the basic and applied level.
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Acknowledgements We are most thankful to James Warbrick-Smith, Angela Douglas and Stephen J. Simpson for their feedback on the manuscript. Many thanks also to Naomi Behmer for her help in finding and ordering obscure references from both the Radcliffe Science Library (Oxford University) and the British Library, and for reading various drafts of the manuscript. This research has been supported by grants from the NSF (MCB 0115401), Asgrow seed company and Monsanto to W.D.N.
References Agarwal, H. C. (1970). Sterol requirements of the beetle Trogoderma. J. Insect Physiol. 16, 2023–2026. Akihisa, T., Shimizu, N., Ghosh, P., Thakur, S., Rosenstein, F. U., Tamura, T. and Matsumoto, T. (1987). Sterols of the Cucurbitaceae. Phytochem. 26, 1693–1700. Akihisa, T., Kokke, W. C. M. C. and Tamura, T. (1991). Naturally occurring sterols and related compounds from plants. In: Physiology and Biochemistry of Sterols (eds. Patterson, G. W., and Nes, W. D.), pp. 172–228. Champaign, IL: American Oil Chemists’ Society. Aldrich, J. R., Kelly, T. J. and Woods, C. W. (1982). Larval moulting hormone of trichophoran Hemiptera-Heteroptera: makisterone A, not 20-hydroxyecdysone. J. Insect Physiol. 28, 857–861. Alexandra, C., Lecourtois, M. and Vincent, J-P. (1999). Wingless and Hedgehog pattern Drosophila denticel belts by regulating the production of short-range signals. Development 126, 5689–5698. Ba, A., Guo, D., Norton, R. A., Phillips, S. A. and Nes, W. D. (1995). Developmental differences in the sterol composition of Solenopsis invicta. Arch. Insect Biochem. Physiol. 29, 1–9. Baker, J. E. (1974). Differential sterol utilization by larvae of Sitophilus oryzae and Sitophilus granarius. Ann. Entomol. Soc. Am. 67, 591–594. Beck, S. D., Lilly, J. H. and Stauffer, J. F. (1949). Nutrition of European corn borer, Pyrausta nubilalis (Hbn.). I. Development of a satisfactory purified diet for larval growth. Ann. Ent. Soc. Am. 42, 483–496. Beenakkers, A. M. T., van der Horst, D. J. and van Marrewijk, W. J. A. (1985). Insect lipids and their role in physiological processes. Prog. Lipid Res. 24, 19–67. Behmer, S. T. and Elias, D. O. (1999a). The nutritional significance of sterol metabolic constraints in the generalist grasshopper Schistocerca americana. J. Insect Physiol. 45, 339–348. Behmer, S. T. and Elias, D. O. (1999b). Phytosterol structure and its impact on feeding behaviour in the generalist grasshopper Schistocerca americana. Physiol. Entomol. 24, 18–27. Behmer, S. T. and Elias, D. O. (2000). Sterol metabolic constraints as a factor contributing to the maintenance of diet mixing in grasshoppers (Orthoptera: Acrididae). Physiol. and Biochem. Zool. 73, 219–230. Behmer, S. T. and Grebenok, R. J. (1998). Impact of sterols on life-history traits of a caterpillar. Physiol. Entomol. 23, 165–175.
60
S. T. BEHMER AND W. D. NES
Behmer, S. T., Elias, D. O. and Bernays, E. A. (1999a). Post-ingestive feedbacks and associative learning regulate the intake of unsuitable sterols in a generalist grasshopper. J. Exp. Biol. 202, 739–748. Behmer, S. T., Elias, D. O. and Grebenok, R. J. (1999b). Phytosterol metabolism and absorption in the generalist grasshopper, Schistocerca americana (Orthoptera: Acridiae). Arch Insect Biochem. Physiol. 42, 13–25. Beilby, J. P. (1980). Fatty acid and sterol composition of ungerminated spores of the vesicular-arbuscular mycorrhizal fungus Acaulospora laevis. Lipids 15, 949–952. Beilby, J. P. and Kidby, D. K. (1980). Sterol composition of ungerminated and germinated spores of the vesicular-arbuscular mycorrhizal fungus, Glomus caledonius. Lipids 15, 375–378. Bergmann, W. (1957). The plant sterols. Ann. Rev. Plant Physiol. 4, 383–426. Bergmann, E. D. and Levinson, Z. H. (1966). Utilization of steroid derivatives by larvae of Musca vicina (Macq.) and Dermestes maculatus Deg. J. Insect Physiol. 12, 77–81. Bernays, E. A. (1992). Plant sterols and host-plant affliliations of herbivores. In: InsectPlant Interactions (ed. Bernays, E. A.), Vol. 4, pp. 45–57. Boca Raton, FL: CRC Press. Billheimer, J. T., Tavani, D. M. and Ritter, K. S. (1983). Acyl coenzyme A: Cholesterol acyltransferase activity in fat body and intestinal microsomes of Heliothis zea. Comp. Biochem. Physiol. 76B, 127–132. Bitmann, R. (1997). Has nature designed the cholesterol side chain for optimal interaction with phospholipids. Subcellular Biochem. 28, 145–171. Bloem, K. A., Kelley, K. C. and Duffey, S. S. (1989). Differential effect of tomatine and its alleviation by cholesterol on larval growth and efficiency of food utilization in Heliothis zea and Spodoptera exigua. J. Chem. Ecol. 15, 387–398. Borror, D. J., Triplehorn, C. A. and Johnson, N. F. (1989). An Introduction to the Study of Insects, 6th edition, p. 875. Orlando, FL: Harcourt Brace & Company. Brown, D. A. and London, E. (1997). Structure of detergent-resistant membrane domains: Does phase separation occur in biological membranes?. Biochem. Biophys. Res. Commun. 240, 1–7. Brust, M. and Fraenkel, G. (1955). The nutritional requirements of the larvae of a blowfly, Phormia regina (Meig.). Physiol. Zool. 28, 186–204. Budowski, P., Ishaaya, I. and Katz, M. (1967). Growth inhibition of Dermestes maculatus by phytosterols. J. Nutrition 91, 201–207. Campbell, B. C. and Nes, W. D. (1983). A reappraisal of sterol biosynthesis and metabolism in aphids. J. Insect Physiol. 29, 149–156. Canavoso, L. E. and Wells, M. A. (2001). Role of lipid transport particle in lipid delivery of diacylglycerol from midgut to lipophorin in larval Manduca sexta. Insect Biochem. Molec. Biol. 31, 783–790. Canavoso, L. E., Yun, H. K., Jouni, Z. E. and Wells, M. A. (2003). Lipid transfer particle mediates the delivery of diacylglycerol from HDLp to fat body in larval Manduca sexta. Insect Biochem. Molec. Biol. (in press). Chambers, P., Sword, G., Angel, J., Behmer, S. and Bernays, E. A. (1996). Foraging by generalist grasshoppers: dietary mixing and the role of crypsis. Animal Behaviour 52, 155–165. Champagne, D. and Bernays, E. A. (1991). Phytosterol unsuitability as a factor mediating food aversion learning in the grasshopper Schistocerca americana. Physiol. Entomol. 16, 391–400. Charlet, M., Roussel, J-P., Rinternecht, E., Berchtold, J-P. and Costet, M-F. (1988). Developmental and morphogenetic alterations in larvae of Locusta migratoria reared on plant diet with a selectively modified sterol profile. J. Insect Physiol. 34, 787–796.
INSECT STEROL NUTRITION AND PHYSIOLOGY
61
Chauvin, R. (1949). Les ste´rols dan le re´gime alimentaire de certains Orthopte`res. C.R. Acad. Sci. 229, 902–903. Chino, H. (1985). Lipid transport: Biochemistry of hemolymph lipophorin. In: Comprehensive Insect Physiology, Biochemistry and Pharmacology (eds. Kerkut, G. A., and Gilbert, L. I.), Vol. 10, pp. 115–135. Oxford: Pergamon Press. Chippendale, G. M. (1971). Lipid requirements of the angoumois grain moth, Sitotroga cerealella. J. Insect Physiol. 17, 2169–2177. Chippendale, G. M. and Reddy, G. P. V. (1972a). Polyunsaturated fatty acid and sterol requirements of the southwestern corn borer, Diatreae grandiosella. J. Insect Physiol. 18, 305–316. Chippendale, G. M. and Reddy, G. P. V. (1972b). Dietary sterols: Role in larval feeding behaviour of the southwestern corn borer, Diatraea grandiosella. Experientia 28, 485–486. Chiu, P-L., Patterson, G. W. and Salt, T. A. (1988). Sterol composition of Pteridophytes. Phytochem. 27, 819–822. Chiu, S. F. and McKay, C. M. (1939). Nutritional studies of the confused flour beetle (Tribolium confusum Duval) and the bean weevil (Acanthoscelides obtectus (Say)). Ann. Entomol. Soc. Am. 32, 164–170. Chu, H., Norris, D. M. and Kok, L. T. (1970). Pupation requirements of the beetle, Xyleborus ferrugineus: Sterols other than cholesterol. J. Insect Physiol. 16, 1379–1387. Clark, A. J. and Bloch, K. (1959a). Function of sterols in Dermetes vulpinus. J. Biol. Chem. 234, 2583–2588. Clark, A. J. and Bloch, K. (1959b). Conversion of ergosterol to 22-dehydrocholesterol in Blatella germanica. J. Biol. Chem. 234, 2589–2594. Clark, E. W. (1973). Hylobius pales: A meridic larval diet using sitosterol. J. Econ. Entomol. 66, 841–843. Clayton, R. B. (1964). The utilization of sterols by insects. J. Lipid Res. 5, 3–19. Clayton, R. B., Hinkle, P. C., Smith, D. A. and Edwards, A. M. (1964). The intestinal absorption of cholesterol, its esters and some related sterols and analogues in the roach, Eurycotis floridana. Comp. Biochem. Physiol. 11, 333–350. Cooke, J. and Sang, J. H. (1970). Utilization of sterols by larvae of Drosophila melanogaster. J. Insect Physiol. 16, 801–812. Cooke, J. and Sang, J. H. (1972). Physiological genetics of melanotic tumours in Drosophila melanogaster, Genet. Res. Camb. 20, 317–329. Corbin, D. R., Grebenok, R. J., Ohnmeiss, T. E., Greenplate, J. T. and Purcell, J. P. (2001). Expression and chloroplast targeting of cholesterol oxidase in transgenic tobacco plants. Plant Physiol. 126, 1116–1128. Costet, M. F., Achouri, M. E., Charlet, M., Lanot, R., Benveniste, P. and Hoffmann, J. A. (1987). Ecdysteroid biosynthesis and embryonic development are disturbed in insects (Locusta migratoria) reared on plant diet (Tritcum sativum) with a selectively modified sterol profile. Proc. Natl. Acad. Sci. (USA) 84, 643–647. Dadd, R. H. (1960). The nutritional requirements of locusts - II. Utilization of sterols. J. Insect Physiol. 5, 161–168. Dadd, R. H. (1977). Qualitative requirements and utilization of nutrients: Insects. In: Handbook Series in Nutrition and Food. Section D: Nutritional Requirements (ed. Rechcigl, M., Jr.), Vol. 1, pp. 305–346. Cleveland, OH: CRC Press. Dadd, R. H. (1985). Nutrition: organisms. In: Comprehensive Insect Physiology, Biochemistry and Pharmacology (eds. Kerkut, G. A., and Gilber, L. I.), Vol. 4, pp. 313–390. Elmsford, NY: Pergamon Press. Dadd, R. H. and Kleinjan, J. E. (1984). Lecithin-dependent phytosterol utilization by larvae of Culex pipiens (Diptera: Culicidae). Ann. Entomol. Soc. Am. 77, 518–525.
62
S. T. BEHMER AND W. D. NES
Dambre-Raes, H. (1976). The effect of dietary cholesterol on the development of Hylemya brassicae. J. Insect Physiol. 22, 1287–1289. Douglas, A. E. (1988). On the source of sterols in the green peach aphid, Myzus persicae, reared on holidic diets. J. Insect Physiol. 34, 403–408. Douglas, A. E. (1998). Nutritional interactions in insect-microbial symbioses: Aphids and their symbiotic bacteria Buchnera. Annu. Rev. Entomol. 43, 17–37. Dupnik, T. D. and Kamm, J. A. (1970). Development of an artificial diet for Crambus trisectus. J. Econ. Entomol. 63, 1578–1581. Dussourd, D. E., Harvis, C. A., Meinwald, J. and Eisner, T. (1991). Pheromonal advertisemet of a nuptial gift by a male moth Utetheisa ornatrix. Proc. Nat. Acad. Sci. (USA) 20, 9224–9227. Dutky, R. C., Robbins, W. E., Kaplanis, J. N. and Shortino, T. J. (1963). The sterol esters of housefly eggs. Comp. Biochem. Physiol. 9, 251–255. Dutky, R. C., Robbins, W. E., Shortino, T. J., Kaplanis, J. N. and Vroman, H. E. (1967). The conversion of cholestanone to cholestanol by the housefly, Musca domestica L. J. Insect Physiol. 13, 1501–1510. Dwivedy, A. K. (1993). A preferential uptake of cholesterol by the brain tissue of the housefly, Musca domestica L. Experientia 39, 272–273. Earle, N. W., Lambremont, N., Burks, M. L., Slatten, B. H. and Bennett, A. F. (1967). Conversion of -sitosterol to cholesterol in the boll weevil and the inhibition of larval development by the two asa sterols. J. Econ. Entomol. 60, 291–293. Erwin, T. L. (1982). Tropical forests: Their richness in Coleoptera and other arthropod species. Coleopterists Bull. 36, 74–75. Feldlaufer, M. F. and Svoboda, J. A. (1991). Sterol utilizaton and ecdysteroid content in the house fly, Musca domestica (L.). Insect Biochem. 21, 53–56. Feldlaufer, M. F., Svoboda, J. A., Aldrich, J. R. and Lusby, W. R. (1986). The neutral sterols of Megalotomus quinquespinosus (Say) (Hemiptera: Alydidae) and identification of makisterone A as the major free ecdysteroid. Arch. Insect Biochem. Physiol. 3, 423–430. Feldlaufer, M. F., Buchmann, S. L., Lusby, W. R., Weirich, G. F. and Svoboda, J. A. (1993). Neutral sterols and ecdysteroids of the solitary cactus bee Diadasia rinconis Cockerell (Hymenoptera: Anthophoridae). Arch. Insect Biochem. Physiol. 23, 91–98. Feldlaufer, M. F., Weirich, G. F., Imberski, R. B. and Svoboda, J. A. (1995). Ecdysteroid production in Drosophila melanogaster reared on defined diets. Insect Biochem. Molec. Biol. 25, 709–712. Feldlaufer, M. F., Harrison, D. J. and Wilzer, K. R. (1997). Comparison of the dietary and tissue sterols of the greater wax moth, Galleria mellonella (L). Apidologie 28, 85–90. Fogleman, J. C., Duperret, S. M. and Kircher, H. W. (1986). The role of phytosterols in host plant utilization by cactophillic Drosophila. Lipids 21, 92–96. Fraenkel, G. and Blewett, M. (1943). The sterol requirements of several insects. Biochem. J. 37, 692–695. Fraenkel, G. and Blewett, M. (1946). The dietics of the clothes moth, Tineola bisselliella Hum. J. Expt. Biol. 22, 156–161. Fraenkel, G., Reid, J. A. and Blewett, M. (1941). The sterol requirements of the larva of the beetle, Dermestes vulpinus Fabr. Biochem. J. 35, 712–720. Galford, J. R. (1972). Some basic nutritional requirements of smaller European elm bark beetle larvae. J. Econ. Entomol. 65, 681–684. Gibson, J. M., Majumder, M. S. I., Mendis, A. H. W. and Rees, H. H. (1983). Absence of phytosterol dealkylation and identification of the major ecdysteroid as
INSECT STEROL NUTRITION AND PHYSIOLOGY
63
makisterone A in Dysdercus fasciatus (Heteroptera, Phrrhocoridae). Arch. Insect Biochem. Physiol. 1, 105–120. Gilbert, L. I., Rybcznski, R. and Warren, J. T. (2002). Control and biochemical nature of the ecdysteroidogenic pathway. Annu. Rev. Ent. 47, 883–916. Gingrich, R. E. (1964). Nutritional studies on screw-worm larvae with chemically defined media. Ann. Entomol. Soc. Am. 57, 351–360. Goldberg, L. and De Meillon, B. (1948). The nutrition of the larva of Ae¨des aegyptii Linnaeus. Biochem. J. 43, 372–377. Goodfellow, R. D., Barnes, F. J. and Graham, W. J., Jr (1971). Dietary sterol effects on mevalonate kinase activity in axenic and non-axenic Sarcophaga bullata (Diptera). J. Insect Physiol. 17, 1625–1635. Gordon, H. T. (1959). Minimal nutritional requirements of the german roach, Blatella germanica L. Ann. N.Y. Acad. Sci. 77, 290–351. Guo, D., Venkatramesh, M. and Nes, W. D. (1995). Developmental regulation of sterol biosynthesis in Zea mays. Lipids 30, 203–219. Guo, D., Jia, Z. and Nes, W. D. (1996). Stereochemistry of hydrogen migration from C-24 to C-25 during phytosterol biomethylation. J. Amer. Chem. Soc. 118, 8507–8508. Hamamura, Y. (1970). The substances that control the feeding behavior and the growth of the silkworm Bombyx mori L. In: Control of Insect Behavior by Natural Products (eds. Wood, D. L., Silverstein, R. M., and Nakajima, M.), pp. 55–80. London: Academic Press. Hamamura, Y., Hayashiya, K., Naito, K., Matsuura, K. and Nishida, J. (1962). Food selection by silkworm larvae. Nature 194, 754–755. Heed, W. B. and Kircher, H. W. (1965). Unique sterol in the ecology and nutrition of Drosophila pachea. Science 149, 758–761. Herbert, E. W., Jr, Svoboda, J. A., Thompson, M. J. and Shimanuki, H. (1980). Sterol utilization in honey bees fed a synthetic diet: Effects on brood rearing. J. Insect Physiol. 26, 287–289. Heywood, V. H. (1993). Flowering Plants of the World. pp. 335. New York: Oxford University Press. Hobson, R. P. (1935a). CCXXXVI. On a fat-soluble growth factor required by blow-fly larvae. II. Identity of the growth factor with cholesterol. Biochem. J. 29, 2023–2026. Hobson, R. P. (1935b). CLVI. On fat-soluble growth factor required by blow-fly larvae. I. distribution and properties. Biochem. J. 29, 1292–1296. House, H. L. (1954). Nutritional studies with Pseudocarcophaga affinis (Fall.), a dipterous parasite of the spruce budworm, Choristoneura fumiferana (Clem.). IV. Effects of ribonucleic acid, glutathione, dextrose, a salt mixture, cholesterol, and fats. Can. J. Zool. 32, 358–365. Hsiao, T. H. and Fraenkel, G. (1968). The influence of nutrient chemicals on the feeding behaviour of the Colorado Potato Beetle, Leptinotarsa decemlineata (Coleoptera: Chrysomelidae). Ann. Ent. Soc. Am. 61, 44–54. Ichimasa, Y. (1976). Sterol accumulation in developing ovaries of the silkworm Bombyx mori in relation to the diapause hormone. J. Insect Physiol. 22, 1071–1074. Ikekawa, N., Morisaki, M. and Fujimoto, Y. (1993). Sterol metabolism in insects: Dealkylation of phytosterol to cholesterol. Acc. Chem. Res. 26, 139–146. Ingham, P. W., Taylor, A. M. and Nakano, Y. (1991). Role of Drosophila patched gene in positional signalling. Nature 353, 184–190. Ishii, S. (1951). Studies on the host-plants of the cowpea weevil (Callosobruchus chinensis L.) IX. On the relation between chemical constitutions of sterols and the development of larva. Botyu Kagaku 16, 83–90.
64
S. T. BEHMER AND W. D. NES
Ishii, S. and Urushibara, H. (1954). On fat and water soluble growth factors required by the rice stem borer, Chilo simplex. Bull. Nat. Inst. Agr. Sci. Japan, Ser. C. 4, 109–133. Ito, T. (1961). Sterol requirements of the silkworm, Bombyx mori. Nature 191, 882–883. Ito, T., Kawashima, K., Nakahara, M., Nakanishi, K. and Terahara, A. (1964). Effects of sterols on feeding and nutrition of the silkworm, Bombyx mori L. J. Insect Physiol. 10, 225–238. Ito, T. and Nakasone, S. (1966). Nutrition of the silkworm, Bombyx mori, XIII. Nutritive effects of fatty acids. Bull. Sericult. Exp. Station Tokyo 21, 385–400. Joshi, M. and Agarwal, H. C. (1976). Cholesterol absorption in the roach, Periplaneta americana. Entomon 1, 93–100. Joshi, M. and Agarwal, H. C. (1977). Site of cholesterol absorption in some insects. J. Insect Physiol. 23, 403–404. Jouni, Z. E., Zamora, J. and Wells, M. A. (2002a). Absorption and tissue distribution of cholesterol in Manduca sexta. Arch. Insect Biochem. Physiol. 49, 167–175. Jouni, Z. E., Yun, H. K. and Wells, M. A. (2002b). Cholesterol efflux from larval Manduca sexta fat body in vitro: High-density lipophorin as the acceptor. J. Insect Physiol. 48, 609–618. Jouni, Z. E., Takada, N., Gazard, J., Maekawa, H., Wells, M. A. and Tsuchida, K. (2003). Transfer of cholesterol and diacylglycerol from lipophorin to Bombyx mori ovarioles in vitro: Role of the lipid transfer particle. Insect Biochem Mol. Biol. 33, 145–153. Kaplanis, J. N., Monroe, R. E., Robbins, W. E. and Louloudes, S. J. (1963). The fate of dietary H3- -sitosterol in the adult house fly. Ann. Entomol. Soc. Am. 56, 198–201. Kaplanis, J. N., Robbins, W. E., Monroe, R. E., Shortino, T. J. and Thompson, M. J. (1965). The utilization and fate of -sitosterol in the larva of the housefly, Musca domestica L. J. Insect Physiol. 11, 251–258. Katz, M., Budowski, P. and Bondi, A. (1971). The effect of phytosterols on the growth and sterol composition of Dermestes maculatus. J. Insect Physiol. 17, 1295– 1303. Keiser, L. S. and Yazlovetski, I. G. (1988). Comparative study of sterols in the larval lacewing Chrysopa carnea and its food species. J. Evol. Biochem. Physiol. 24, 108–113. Kircher, H. W. (1982). Sterols and insects. In: Cholesterol Systems in Insects and Animals (ed. Dupont, J.), pp. 1–50. Boca Raton, FL: CRC Press. Kircher, H. W., Rosenstein, F. U. and Fogleman, J. C. (1984). Selective uptake and lack of dealkylation of phytosterols by cactophilic species of Drosophila. Lipids 19, 235–238. Komnick, H. and Giesa, U. (1994). Intestinal absorption of cholesterol, transport in the haemolymph, and incorporation into the fat body and malpighian tubules of the larval dragonfly Aeshna cyanea. Comp. Biochem. Physiol. 107A, 553–557. Krebs, K. C. and Lan, Q. (2003). Isolation and expression of a sterol carrier protein-2 gene from the yellow fever mosquito, Aedes aegypti. Insect Mol. Biol. 12, 51–60. Kristensen, N. P. (1991). Phylogeny of extant arthropods. In: Insects of Australia: A Textbook for Students and Research Workers, 2nd ed., Vol. 1, (eds. Naumann, I. D., Carne, P. B., Lawrence, J. F., Nielsen, E. S., Spradberry, J. P., Taylor, R. W., Whitten, M. J., and Littlejohn, M. J.), pp. 125–140. Carlton, Victoria: Melbourne University Press. Kuthiala, A. and Ritter, K. S. (1988). Esterification of cholesterol and cholestanol in the whole body, tissue, and frass of Heliothis zea. Arch. Insect Biochem. Physiol. 7, 237–247.
INSECT STEROL NUTRITION AND PHYSIOLOGY
65
Lasser, N. L. and Clayton, R. B. (1966). The intracellular distribution of sterols in Eurycotis floridana and its possible relation to subcellular membrane structure. J. Lipid Res. 7, 413–421. Lasser, N. L., Edwards, A. M. and Clayton, R. B. (1966). Distribution and dynamic state of sterols and steroids in the tissues of an insect, the roach Eurycotis floridana. J. Lipid Res. 7, 403–411. Leclercq, J. (1948). Sur les besoins en ste´rols des larves de Tenebrio molitor L. Biochim. Biophys. Acta 2, 614–617. Lee, J. C. and Bernays, E. A. (1988). Declining acceptability of a food plant for the polyphagous grasshopper Schistocerca americana: The role of food aversion learning. Physiol. Entomol. 13, 291–301. Lehrer, A. T., Dugassa-Gobena, D., Vidal, S. and Seifert, K. (2000). Transport of resistance-inducing sterols in phloem sap of barley, Z. Naturforsch. 55C, 948–952. Levinson, Z. H. (1962). The function of dietary sterols in phytophagous insects. J. Insect Physiol. 8, 191–198. Levinson, Z. H. and Gothilf, S. (1955). A semisynthetic diet for axenic growth of the carob moth Ectomyelois ceratoniae (Zell.). Riv. Parassit. 26, 19–26. Levinson, Z. H. and Bergmann, E. D. (1957). Steroid utilization and fatty acid synthesis by the larva of the housefly Musca vicina Macq. Biochem. J. 65, 254–260. Lubzens, E., Tietz, A., Pines, M. and Applebaum, S. W. (1981). Lipid accumulation in oocytes of Locusta migratoria migratorioides. Insect Biochem. 11, 323–329. Lusby, W. R., Buchmann, S. L. and Feldlaufer, M. F. (1993). Pollen sterols from three species of Sonoran cacti. Lipids 28, 469–470. Lutjohann, D., Bjorkhem, I., Beil, U. F. and von Bergmann, K. (1995). Sterol absorption and sterol balance in phytosterolemia evaluated by deuterium-labeled sterols: Effects of sitostanol treatment. J. Lipid Res. 36, 1763–1773. Macauley, S. K., Billheimer, J. T. and Ritter, K. S. (1986). Sterol substrate specificity of acyl coenzyme A: Cholesterol transferase from the corn earworm, Heliothis zea. J. Lipid Res. 27, 64–71. MacDonald, D. L. and Ritter, K. S. (1988). Effect of host sterols on the sterol composition and virulence of a nuclear polyhedrosis virus of Heliothis Zea. Lipids 23, 1107–1113. MacDonald, D. L., Nham, D. N., Cochran, W. K. and Ritter, K. S. (1990). Differences in the sterol composition of Heliothis zea fed Zea mays versus Medicago satvia. Insect Biochem. 20, 437–442. Maddison, D. R., Maddison, W. P., Schulz, K.-S., Wheeler, T. and Frumkin, J. (2001). The Tree of Life Web Project. Internet address: http://tolweb.org. Magis, N. (1954). Les besions nutritifs des larves de Tribolium confusum Duv. (Coleoptera Tenebrionidae). Bull. Ann. Soc. Entomol. Belg. 90, 49–58. Mangla, A. T. and Nes, W. D. (2000). Sterol C-methyl transferase from Prototheca wickerhamii. Mechanism, sterol specificity and inhibition. Bioorg. Med. Chem. 8, 925–936. Maurer, P., Royer, C., Mauchamp, B., Porcheron, P., Debieu, D. and Riba, G. (1991). Occurrence of 28- and 27-carbon ecdysteroids and sterols in developing worker pupae of the leaf-cutting ant Acromyrmex octospinosus (Reich) (Hymenoptera, Formicidae: Attini). Arch. Insect Biochem. Physiol. 16, 1–9. Maurer, P., Debieu, D., Malosse, C., Leroux, P. and Riba, G. (1992). Sterols and symbiosis in the leaf-cutting ant Acromyrmex octospinosus (Reich) (Hymenoptera, Formicidae: Attini). Arch. Insect Biochem. Physiol. 20, 13–21. Maurer, P., Girault, J.-P., Larcheveque, M. and Lafont, R. (1993). 24-epi-makisteroneA (not makisterone-A) is the major ecdysteroid in the leaf-cutting ant Acromyrmex
66
S. T. BEHMER AND W. D. NES
octospinosus (Reich) (Hymenoptera, Formicidae: Attini). Arch. Insect Biochem. Physiol. 23, 29–35. McKennis, H. (1947). The growth-promoting activity of cholesterol and related compounds in black carpet beetle larvae. J. Biol. Chem. 167, 645–654. Mondy, N. and Corio-Costet, M. (2000). The response of the grape berry moth (Lobesia botrana) to a dietary phytopathogenic fungus (Botrytis cinerea): The significance of fungus sterols. J. Insect Physiol. 46, 1557–1564. Monroe, R. E. (1959). Role of cholesterol in house fly reproduction. Nature 184, 1513. Monroe, R. E., Kaplanis, J. N. and Robbins, W. E. (1961). Sterol storage and reproduction in the house fly. Ann. Entomol. Soc. Am. 54, 537–539. Myant, N. B. (1981). The Biology of Cholesterol and Related Steroids. pp. 910. London: William Heinemann Medical Books Ltd. Nayar, J. K. and Fraenkel, G. (1962). The chemical basis of hostplant selection in the silkworm, Bombyx mori (L.). J. Insect Physiol. 8, 505–525. Nes, W. D. (1987). Metabolism and function of sterols in Phytophthora cactorum. J. Am. Oil Chem. Soc. 64, 634–634. Nes, W. D. (1990). Control of sterol biosynthesis and its importance to developmental regulation and evolution. Rec. Adv. Phytochem. 24, 283–327. Nes, W. D. and Lee, P. H. (1990). Evidence for separate intermediates in the biosynthesis of 24b-methylsterol end products by Gibberella fujikuroi. Biochim. Biophys. Acta, 1042, 119–125. Nes, W. D. and Schmidt, J. O. (1988). Isolation of 25(27)-dehydrolanost- -enol from Cerens gigantens and its biosynthetic implications. Phytochem. 27, 1705–1708. Nes, W. D., Xu, S. and Haddon, W. F. (1988a). Evidence for similarities and differences in the biosynthesis of fungal sterols. Steroids 53, 533–558. Nes, W. D., Norton, R. A., Parish, E. J., Meenan, A. and Popja´k, G. (1988b). Concerning the role of 24,25-dihydrolanosterol and lanostanol in sterol biosynthesis by cultured cells. Steroids 53, 461–475. Nes, W. D., Norton, R. A. and Benson, M. (1992). Carbon-13 NMR studies on sitosterol biosynthesized from [13C]mevalonates. Phytochem. 31, 811–811. Nes, W. D., Lopez, M., Zhou, W., Guo, D., Dowd, P. F. and Norton, R. A. (1997). Sterol utilization and metabolism by Heliothis zea. Lipids 32, 1317–1323. Nes, W. D., Marshall, J. A., Zhou, W., Le, H. and Dennis, A. L. (1998). Mechanismbased active site modification of sterol methyl transferase by tritium-labeled 26-homocholesta-8-14-24-trein-26-yn-3 -ol. Tetrahedron Lett. 39, 8575–8578. Nes, W. R. and Nes, W. D. (1980). Lipids in Evolution. pp. 244. New York: Plenum Press. Nes, W. R. and McKean, M. L. (1977). Biochemistry of Steroids and Other Isopentenoids. pp. 690. Baltimore, MD: University Park Press. Nes, W. R., Sekula, B. C., Nes, W. D. and Adler, J. H. (1978). The functional importance of structural features of ergosterol in yeast. J. Biol. Chem. 253, 6218–6225. Nes, W. R., Adler, J. H., Billheimer, J. T., Erickson, K. A., Joseph, J. M., Landrey, J. R., Maraccio-Joseph, R., Ritter, K. S. and Conner, R. L. (1982). A comparison of the biological properties of androst-5-en-3 -ol, a series of (20R)-n-alkylpregn5-en-3 -ols and 21-isopentylcholesterol with those of cholesterol. Lipids 17, 257–262. Noda, H. and Koizumi, Y. (2003). Sterol biosynthesis by symbionts: cytochrome P450 sterol C-22 desaturase genes from yeastlike symbiotes of rice planthoppers and anobiid beetles. Insect Biochem. Mol. Biol. 33, 649–658. Noland, J. L. (1954). Sterol metabolism in insects. I. Utilization of cholesterol derivatives by the cockroach Blattella germanica L. Arch. Biochem. Biophys. 48, 370–379.
INSECT STEROL NUTRITION AND PHYSIOLOGY
67
Norris, D. M., Baker, J. M. and Chu, H. M. (1969). Symbiotic interrelationships between microbes and ambrosia beetles. III. Ergosterol as the source of sterol to the insect. Ann. Entomol. Soc. Am. 62, 413–414. Ostlund, R. E., Jr. (2002). Phytosterols in human nutrition. Annu. Rev. Nutr. 22, 533–549. Ostlund, R. E., Jr, McGill, J., Zeng, C-M, Covey, D. F. and Stearns, J. (2002). Gastrointestinal absorption and plasma kinetics of soy 5-phytosterols and phytostanols in humans. Am. J. Physiol. Endocrinol. Metab. 282, E911–E916. Parker, S. R. and Nes, W. D. (1992). Regulation of sterol biosynthesis and its phylogenetic implications. ACS Symp. Ser. 497, 110–145. Patterson, G. W. (1994). Phylogenetic distribution of sterols. In: Isopentenoids and Other Natural Products: Evolution and Function (ed. Nes, W. D.), pp. 91–107. Washington DC: American Chemical Society. Pausch, R. D. and Fraenkel, G. (1966). The nutrition of the larva of the oriental rat flea Xenopsylla cheopis (Rothschild). Physiol. Zool. 39, 202–222. Popja´k, G., Edmond, J., Anet, F. A. L. and Easton, N. R., Jr (1977). Carbon-13 NMR studies on cholesterol biosynthesized from [13C]mevalonates. J. Amer. Chem. Soc. 99, 931–935. Porter, J. A., Young, K. E. and Beachy, P. A. (1996). Cholesterol modification of hedgehog signaling proteins in animal development. Science 274, 255–259. Prestwich, G. D., Shieh, H. M. and Gayen, A. K. (1983). Side chain modified sterols as probes into insect molting hormone metabolism. II. Synthesis of monofluorocholesterols. Steroids 41, 79–94. Prestwich, G. D., Yamaoka, R., Phirwa, S. and Depalma, A. (1984). Isolation of 2fluorocitrate produced by in vivo dealkyation of 29-fluorostigmasterol in an insect. J. Biol. Chem. 259, 1022–1026. Rao, A. V. and Korathkar, R. (1997). Anticarcinogenic effects of saponins and phytosterols. In: ACS Symposium Series 662: Antinurients and Phytochemicals in Food (ed. Shahidi, F.), pp. 313–324. Washington DC: American Chemical Society. Rasmussen, S. (1956). On the significance of cholesterol and yeast extract in the diet of larvae of house longhorn beetle (Hylotrupes bajulus). Oikos 7, 243–250. Rasmussen, S. (1958). On the response of Hylotrupes bajulus larvae to doses of cholesterol and other sterols. Oikos 9, 211–220. Rath, R., Singhal, R., Goel, V. and Agarwal, H. C. (1993). In vivo and in vitro metabolism of sitosterol in Heliothis armigera (Hubner) and Locusta migratoria L. Indian J. Exp. Biol. 31, 699–703. Richmond, J. A. and Thomas, H. A. (1975). Hylobius pales: Effect of dietary sterols on development and on sterol content of somatic tissue. Ann. Entomol. Soc. Am. 68, 329–332. Rietveld, A., Neutz, S., Simons, K. and Eaton, S. (1999). Association of sterol- and glycolphosphatidylinositol-linked proteins with Drosophila raft lipid microdomains. J. Biol. Chem. 274, 12049–12054. Ritter, K. S. (1984). Metabolism of 0-, 5-, and 7-sterols by larvae of Heliothis zea. Arch. Insect Biochem Physiol. 1, 281–296. Ritter, K. S. (1986). Utilization of 5,7- and 8-sterols by larvae of Heliothis zea. Arch. Insect Biochem. Physiol. 3, 349–362. Ritter, K. S. (1988). Steinernema feltiae ( ¼ Neoaplectana carpocapsae): effects of sterols and hypolipidemic agents on development. Experimental Parasitology 67, 257–267. Ritter, K. S. and Johnson, J. A. (1991). Effects of host sterols on the development and sterol composition of Microplitis demolitor (Hymenoptera: Braconidae) in Heliothis zea (Lepidoptera: Noctuidae). Ann. Entomol. Soc. Am. 84, 79–86.
68
S. T. BEHMER AND W. D. NES
Ritter, K. S. and Nes, W. R. (1981a). The effects of cholesterol on the development of Heliothis zea. J. Insect Physiol. 27, 175–182. Ritter, K. S. and Nes, W. R. (1981b). The effects of the structure of sterols on the development of Heliothis zea. J. Insect Physiol. 27, 419–424. Ritter, K. S., Weiss, B. A., Norrbom, A. L. and Nes, W. R. (1982). Identification of 5,7-24-methylene- and methylsterols in the brain and whole body of Atta cephalotes isthmicola. Comp. Biochem. Physiol. 71B, 345–349. Robbins, W. E. (1963). Studies on the utilization, metabolism and function of sterols in the housefly, Musca domestica. In: Radiation and Radioisotopes Applied to Insects of Agricultural Importance, pp. 269–280. Proceedings of a Symposium, Athens, 22–26 April 1963. Vienna: International Atomic Energy Agency. Robbins, W. E., Dutky, R. C., Monroe, R. F. and Kaplanis, J. N. (1962). The metabolism of H3- -sitosterol by the German cockroach. Ann. Entomol. Soc. Am. 55, 102–104. Robbins, W. E., Kaplanis, J. N., Svoboda, J. A. and Thompson, M. J. (1971). Steroid metabolism in insects. Ann. Rev. Ent. 16, 53–72. Rock, G. C. (1969). Sterol and water-soluble vitamin requirements for larvae of the red-banded leaf roller, Argyrotaenia velutinana. Ann. Entomol. Soc. Am. 62, 611–613. Rowell, C. H. F. and Flook, P. K. (1998). Phylogeny of the Califera and the Orthoptera as derived from ribosomal gene sequences. J. Orthop. Res. 7, 147–156. Ryan, R. O. and van der Horst, D. J. (2000). Lipid transport biochemistry and its role in energy production. Annu. Rev. Ent. 45, 233–260. Rzama, A., Dufourc, E. J. and Arreguy, B. (1994). Sterols from green and blue-green algae grown on reused waste water. Phytochem. 37, 1625–1628. Sall, C., Tsoupras, G., Kappler, C., Lagueux, M., Zachary, D., Luu, B. and Hoffman, J. A. (1983). Fate of maternal conjugated ecdysteroids during embryonic development in Locusta migratoria. J. Insect Physiol. 29, 491–507. Salt, T. A., Sihua, X., Patterson, G. W. and Adler, J. H. (1991). Diversity of sterol biosynthetic capacity in the Caryophyllidae. Lipids 26, 604–613. Sarma, P. S. and Sreenivasaya, M. (1941). Studies in insect nutrition: The nature of the fat-soluble factor. Current Sci. (India) 10, 525–526. Schaefer, C. H., Kaplanis, J. N. and Robbins, W. E. (1965). The relationship of the sterols of the Virginia pine sawfly, Neodiprion pratti Dyar, to those of two host plants, Pinus virginiana Mill and Pinus rigida Mill. J. Insect Physiol. 11, 1013–1021. Schiff, N. M. and Feldlaufer, M. F. (1996). Neutral sterols of sawflies (Symphyta): Their relationship to other Hymenoptera. Lipids 31, 441–443. Sculley, C. E. and Boggs, C. L. (1996). Mating systems and sexual division of foraging effort affect puddling behaviour by butterflies. Ecol. Entomol. 21, 193–197. Seckbach, J. S., Ikan, R., Ringelberg, D. and White, D. (1993). Sterols and phylogeny of the acidophilic hot springs algae Cyanidium caldarium and Galdieria sulphuraria. Phytochem. 34, 1345–1349. Sedee, P. D. J. W. (1961). Intermediary metabolism in aseptically reared blowfly larvae, Calliphora erythrocephala (Meig.). I. Biosynthesis of squalene and cholesterol. Arch. Intern. Physiol. Biochim. 69, 284–294. Sehgal, S. S. and Agarwal, H. C. (1973). Quantitative dietary requirements of the beetle Trogoderma for cholesterol and choline. J. Insect Phsyiol. 19, 419–425. Shanks, C. H., Jr and Doss, R. P. (1987). Feeding responses by adults of five species of weevils (Coleoptera: Curculionidae) to sucrose and sterols. Ann. Entomol. Soc. Am. 80, 41–46.
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Short, J. A., Guo, D-A., Svoboda, J. A. and Nes, W. D. (1996). Mechanistic and metabolic studies of sterol 24,25-double reduction in Manduca sexta. Arch. Insect Biochem. Physiol. 31, 1–22. Silberkang, M., Havel, C. M., Friend, D. S., McCarthy, B. J. and Watson, J. A. (1983). Isoprene synthesis in isolated embryonic Drosophila cells. I. Sterol deficient eukaryotic cells. J. Biol. Chem. 258, 8503–8511. Silverman, P. H. and Levinson, Z. H. (1954). Lipid requirements of the larva of the housefly Musca vicina (Macq.) reared under non-aseptic conditions. Biochem. J. 58, 291–294. Simons, K. and Ikonen, E. (1997). Functional rafts in cell membranes. Nature 387, 569–572. Sivapalan, P. and Gnanapragasam, N. C. (1979). The influence of linoleic acid and linolenic acid on adult moth emergence of Homona coffearia from meridic diets in vitro. J. Insect Physiol. 25, 393–398. Soulages, J. L. and Wells, M. A. (1994). Lipophorin: The structure of an insect lipoprotein and its role in lipid transport in insects. Adv. Protein Chem. 45, 371–415. Southwood, T. R. E. (1973). The insect/plant relationship – an evolutionary perspective. In: Insect/Plant Relationships-R.E.S. Symposium 6 (ed. van Emden, H. F.), pp. 3–30. Oxford: Blackwell Scientific Publications. Speight, M. R., Hunter, M. D. and Watt, A. D. (1999). Ecology of Insects: Concepts and Applications. pp. 350. Oxford: Blackwell Science Ltd. Standifer, L. N., Devys, M. and Barbier, M. (1968). Pollen sterols – a mass spectrographic survey. Phytochem. 7, 1361–1365. Strong, D. R., Lawton, J. H. and Southwood, T. R. E. (1984). Insects on Plants: Community Patterns and Mechanisms. pp. 313. Cambridge, MA: Harvard University Press. Svoboda, J. A. (1984). Insect steroids: Metabolism and function. In: Isopentenoids in Plants: Biochemistry and Function (eds. Nes, W. D., Fuller, G., and Tsai, L-S.), pp. 367–388. New York: Dekker. Svoboda, J. A. (1994). Sterol metabolism as a target for insect control. Biochem. Soc. Trans. 22, 635–641. Svoboda, J. A. (1999). Variability of metabolism and function of sterols in insects. Crit. Rev. Biochem. Mol. Biol. 34, 49–57. Svoboda, J. A. and Feldlaufer, M. F. (1991). Neutral sterol metabolism in insects. Lipids 26, 614–618. Svoboda, J. A. and Lusby, W. R. (1986). Sterols of phytophagous and Omnivorous species of Hymenoptera. Arch. Insect Biochem. Physiol. 3, 13–18. Svoboda, J. A. and Lusby, W. R. (1994). Variability of sterol utilization in storedproducts insects. Experientia 50, 72–74. Svoboda, J. A. and Robbins, W. E. (1967). Conversion of -sitosterol to cholesterol blocked in an insect by hypocholesterolemic agents, Science. Wash. 156, 1637–1638. Svoboda, J. A. and Robbins, W. E. (1968). Desmosterol as a common intermediate in the conversion of a number of C28 and C29 plant sterols to cholesterol by the tobacco hornworm. Experientia 24, 1131–1132. Svoboda, J. A. and Robbins, W. E. (1971). The inhibitive effects of azasterols on sterol metabolism and growth and development in insects with special reference to the tobacco hornworm. Lipids 6, 113–119. Svoboda, J. A. and Robbins, W. E. (1978). Comparison of sterols from a phytophagous and predacious species of the family Coccinellidae. Experientia 35, 186–187. Svoboda, J. A. and Thompson, M. J. (1973). Unusual composition of sterols in a phytophagous insect, Mexican bean beetle reared on soybean plants. Lipids 9, 752–755.
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Svoboda, J. A. and Thompson, M. J. (1985). Steroids. In: Comprehensive Insect Physiology, Biochemistry and Pharmacology (eds. Kerkut, G. A., and Gilbert, L. I.), Vol. 10, pp. 137–175. Elmsford, NY: Pergamon Press. Svoboda, J. A. and Weirich, G. F. (1995). Sterol metabolism in the tobacco hornworm, Manduca sexta – a review. Lipids 30, 263–267. Svoboda, J. A., Dutky, S. R., Robbins, W. E. and Kaplanis, J. N. (1976). Sterol composition and phytosterol utilization and metabolism in the milkweed bug. Lipids 12, 318–321. Svoboda, J. A., Thompson, M. J., Robbins, W. E. and Kaplanis, J. N. (1978). Insect steroid metabolism. Lipids 13, 742–753. Svoboda, J. A., Nair, A. M. G., Agarwal, N., Agarwal, H. C. and Robbins, W. E. (1980a). The sterols of the kharpa beetle, Trogoderma granarium Everts. Experientia 35, 1454–1455. Svoboda, J. A., Nair, A. M. G., Agarwal, N. and Robbins, W. E. (1980b). Lack of conversion of C29-phytosterols to cholesterol in the kharpa beetle, Trogoderma granarium Everts. Experientia 36, 1029–1030. Svoboda, J. A., Thompson, M. J., Herbert, E. W., Shortino, T. J. and Szczepanik-van Eeuwin, P. A. (1982). Utilization and metabolism of dietary sterols in the honey bee and yellow fever mosquito. Lipids 17, 220–225. Svoboda, J. A., Herbert, E. W., Jr, Lusby, W. R. and Thompson, M. J. (1983). Comparison of sterols of pollens, honeybee workers, and prepupae from field sites. Arch. Insect Biochem Physiol. 1, 25–31. Svoboda, J. A., Lusby, W. R. and Aldrich, J. R. (1984). Neutral sterols of representatives of two groups of Hemiptera and their correlation to ecdysteroid content. Arch. Insect Biochem. Physiol. 1, 139–145. Svoboda, J. A., Herbert, E. W., Jr, Thompson, M. J. and Feldlaufer, M. F. (1986). Selective sterol transfer in the honey bee: its significance and relationship to other hymenoptera. Lipids 21, 97–101. Svoboda, J. A., Imberski, R. B. and Lusby, W. R. (1989a). Drosophila melanogaster does not dealkylate [14C]sitosterol. Experientia 45, 983–985. Svoboda, J. A., Rees, H. H., Thompson, M. J. and Hoggard, N. (1989b). Intermediates of stigmasterol metabolism in Spodoptera littoralis. Steroids 53, 329–343. Svoboda, J. A., Feldhaufer, M. F. and Weirich, G. F. (1994). Evolutionary aspects of steroid utilization in insects. ACS Symp. Ser. 562, 126–139. Svoboda, J. A., Ross, S. A. and Nes, W. D. (1995a). Comparative studies of metabolism of 4-desmethyl, 4-monomethyl and 4,4-dimethyl sterols in Manduca sexta. Lipids 30, 91–94. Svoboda, J. A., Schiff, N. M. and Feldlaufer, M. F. (1995b). Sterol composition of three species of sawflies (Hymenoptera: Symphyta) and their dietary plant material. Experientia 51, 150–152. Thompson, M. J. (1984). Contributions of USDA scientists to the chemistry and biochemistry of steroids and other isopentenoids. In: Isopentenoids in Plants: Biochemistry and Function (eds. Nes, W. D., Fuller, G., and Tsai, L-S.), pp. 1–27. New York: Dekker. Thompson, M. J., Kaplanis, J. N., Robbins, W. E. and Svoboda, J. A. (1973). Metabolism of steroids in Insects. Adv. Lipid Res. 11, 219–265. Thompson, S. N. (1981). Brachymeria lasus and Pachycrepoideus vindemiae: sterol requirements during larval growth of two hymenopterous insect parasites reared in vitro on chemically defined media. Expt. Para. 51, 220–235. Tint, G. S., Batta, A. K., Xu, G., Shefer, S., Honda, A., Irons, M., Elias, R. and Salen, G. (1997). The Smith-Lemli-Opitz syndrome: A potentially fatal birth defect
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71
caused by a block in the last enzymatic step in cholesterol biosynthesis. Subcellular Biochem. 28, 117–144. Tsuchida, K. and Wells, M. A. (1990). Isolation and characterization of a lipoprotein receptor from the fat body of an insect. J. Biol. Chem. 265, 5761–5767. Turunen, S. (1985). Absorption. In: Comprehensive Insect Physiology, Biochemistry and Pharmacology (eds. Kerkut, G. A., and Gilber, L. I.), Vol. 4, pp. 241–277. Elmsford, NY: Pergamon Press. Turunen, S. and Chippendale, G. M. (1977). Lipid absorption and transport: sectional analysis of the larval midgut of the corn borer, Diatraea grandiosella. Insect Biochem. 7, 203–208. Turunen, S. and Crailsheim, K. (1996). Lipid in sugar absorption. In: Biology of the Insect Midgut (eds. Lehane, M. J., and Billingsley, P. F.), pp. 291–320. London: Chapman and Hall. van Eijk, G. W. and Roeymans, H. J. (1982). Distribution of carotenoids and sterols in relation to the taxonomy of Taphrina and Protomyces. A. van Leeuw. J. Microbiol. 48, 257–264. Vanderzant, E. S. (1963). Nutrition of the boll weevil larva. J. Econ. Entomol. 56, 357–362. Vanderzant, E. S. and Reiser, R. (1956). Studies of the nutrition of the pink bollworm using purified casein media. J. Econ. Entomol. 49, 454–458. Volkman, J. K. (2003). Sterols in microorganisms. Appl. Microbiol. Biotechnol. 60, 495–506. Vroman, H. E., Kaplanis, J. N. and Robbins, W. E. (1964). Cholesterol turnover in the American cockroach, Periplaneta americana (L.). J. Lipid Res. 5, 418–421. Ward, G. W., Mayer, R. T., Feldlaufer, M. F. and Svoboda, J. A. (1991). Gut chitin synthase and sterols from larvae of Diaprepes abbreviatus (Coleoptera: Curculionidae). Arch. Insect Biochem. Physiol. 18, 105–117. Weber, N. A. (1972). Gardening ants: The Attines. Mem. Am. Philos. Soc. 92, 1–146. Weete, J. D. (1973). Sterols of the fungi: Distribution and biosynthesis. Phytochem. 12, 1843–1864. Weete, J. D. and Laseter, J. L. (1974). Distribution of sterols in the fungi 1. Fungal spores. Lipids 9, 575–581. Weete, J. D., Kulifaj, M., Montant, C., Nes, W. R. and Sancholle, M. (1985). Distribution of sterols in fungi. 2. Brassicasterol in tuber and Terfezia species. Can. J. Microbiol. 31, 1127–1130. Weissenberg, M., Levy, A., Svoboda, J. A. and Ishaaya, I. (1998). The effect of some Solanum steroidal alkaloids and glycoalkaloids on larvae of the red flour beetle, Tribolium castaneum, and the tobacco hornworm, Manduca sexta. Phytochem. 47, 203–209. Wensler, R. J. and Dudzinski, A. E. (1972). Gustation of sugars, amino acids and lipids by larvae of the Scarabeid, Sericesthis geminata (Coleoptera). Ent. Exp. Appl. 15, 155–165. Wiklund, C., Kaitala, A., Lindorfs, V. and Abenius, J. (1993). Polyandry and its effect on female reproduction in the green-veined white butterfly Pieris napi (L.). Behav. Ecol. Sociobiol. 33, 25–33. Wood, T. G. and Thomas, R. J. (1973). The mutalistic association between Macrotermitinae and Termitomyces. In: Insect-Fungus Interactions (eds. Wilding, N., Collins, N. M., Hammond, P. M., and Weber, J. F.), pp. 69–92. San Diego: Academic Press. Xu, S., Norton, R. A., Crumley, F. G. and Nes, W. D. (1988). Comparison of the chromatographic properties of sterols, select additional steroids and triterpenoids: Gravity flow liquid chromatography, thin layer chromatography, gas-liquid
72
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chromatography and high-performance liquid chromatography. J. Chromatogr. 452, 377–389. Yamada, J., Morisaki, M., Iwai, K., Hamada, H., Sato, N. and Fujimoto, Y. (1997). 24-Methyl and 24-Ethyl-24(25)-cholesterols as immediate biosynthetic precursors of 24-alkyl sterols in higher plants. Tetrahedron 53, 877–884. Yun, H. K., Jouni, Z. E. and Wells, M. A. (2002). Characterization of cholesterol transport from midgut to fat body in Manduca sexta larvae. Insect Biochem. Molec. Biol. 32, 1151–1158. Zhuang, M., Oltean, D. I., Go´mez, I., Pullikuth, A. K., Sobero´n, M., Bravo, A. and Gill, S. S. (2002). Heliothis virescens and Manduca sexta lipid rafts are involved in Cry1A toxin binding to the midgut epithelium and subsequent pore formation. J. Biol. Chem. 277, 13863–13872.
The Nutritional Physiology of Aphids Angela E. Douglas Department of Biology, University of York, PO Box 373, York, YO10 5YW, UK
1 Introduction 74 2 The diet of aphids: phloem sap 76 2.1 Phloem sap transport 76 2.2 The composition of phloem sap 79 2.3 Aphid-induced changes to phloem sap 85 3 Acquisition of phloem sap by aphids 87 3.1 Ingestion of phloem sap 87 3.2 Stylet penetration to the sieve elements 90 3.3 Aphid saliva 91 4 Processing of food in the aphid alimentary tract 92 4.1 Approaches to study food processing 93 4.2 Structural organization of the alimentary tract 94 4.3 Processing of ingested sugars 97 4.4 Processing of ingested nitrogenous compounds 102 4.5 The fate of ingested allelochemicals 104 5 Acquisition of nutrients from symbiotic micro-organisms 107 5.1 The microbiology of aphids 107 5.2 The nutritional contribution of Buchnera to aphids 109 6 Fate of nutrients acquired by aphids 112 6.1 Metabolic fate of acquired nutrients 112 6.2 Physiological fate of acquired nutrients 117 6.3 Determinants of nutrient allocation patterns 120 7 Future directions 121 7.1 Regulation of nutrient utilization 122 7.2 Comparative physiology of phloem sap feeders 125 8 Concluding comments: why study the nutritional physiology of phloem sap feeding insects 128 References 130
Abstract Plant phloem sap, the principal food of aphids, is a grossly unbalanced diet for animals, with high ratios of sugars:amino acids, non-essential:essential amino acids and K þ :Na þ and exceptionally low lipid levels. The chief digestive ADVANCES IN INSECT PHYSIOLOGY VOL. 31 ISBN 0-12-024231-1 DOI: 10.1016/S0065-2806(03)31002-1
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function of the alimentary tract is hydrolysis of phloem sugars, usually sucrose to glucose and fructose. Additionally, multiple complex sugars, including oligosaccharides of 20 hexose units are synthesized in the gut and these transformations contribute to the reduction in osmotic pressure of ingested phloem sap, such that the voided honeydew is isosmotic with the haemolymph. Sugars assimilated into aphid tissues are allocated principally to respiration and lipid synthesis. Aphids derive their nitrogen requirement from the phloem amino acids and symbiotic bacteria, Buchnera, which provide essential amino acids. Buchnera may also contribute to aphid nitrogen nutrition by recycling aphid waste nitrogen. Recent advances in analytical techniques and molecular biology/genomics provide the basis to explore the processes by which nutrient provisioning by Buchnera is integrated into the wider nutritional physiology of aphids and to identify the physiological traits of Hemiptera which contribute to the predisposition of these insects to evolve the phloem sap feeding habit.
1
Introduction
Insect nutritional physiology addresses the processing of ingested food and pattern of nutrient allocation in an insect. This subject seeks mechanistic explanations of physiological patterns in terms of the activity of specific digestive enzymes or gut transporters and the metabolic characteristics of specific cells or organs; and it offers mechanistic explanations for wholeorganism traits of ecological or evolutionary importance, such as food preferences, habitat requirements or life history traits of the insect. The entomological focus of this review is aphids, interpreted here as the family Aphididae (Blackman and Eastop, 1984). These insects feed principally or exclusively on the phloem sap of plants, and the nutritional physiology of phloem sap feeding insects is predicted to have unusual, possibly unique, characteristics for two linked reasons. The exceptional features of their diet The phloem sap passing more-or-less continuously through the gut of an aphid is a liquid of high osmotic pressure containing readily assimilated nutrients of a composition that is nutritionally grossly unbalanced for animals. It is only by a remarkable array of anatomical, physiological and biochemical adaptations that aphids neither shrivel, through osmotic collapse, as they feed nor die prematurely through malnutrition. The intimate symbiosis with micro-organisms All insect groups using plant sap as the principal or sole source of food have symbiotic micro-organisms, and most aphids bear bacteria of the genus Buchnera, from which they derive nutrients in short supply in phloem sap. When aphids are experimentally
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deprived of their bacteria, for example by antibiotic treatment, they grow poorly and produce no viable offspring. Understanding the physiological characteristics that mediate the successful utilization of phloem sap by the aphid–bacterial symbiosis is not only of academic interest. Aphids include a number of pests of agricultural importance, especially in temperate and glasshouse crops, by both transmission of plant viruses and direct damage. Unusual or unique features of these insects are potential targets for novel approaches to control aphid pests. Although aphids are just one of several hemipteran groups which subsist principally on phloem sap (Fig. 1), they have been the focus of most physiological research on phloem-feeders. Why aphids? The principal reason is that some aphid species can be cultured very easily under laboratory conditions. The cultured aphids are parthenogenetic, viviparous females, which, although small, have a remarkable reproductive capacity. For example, an adult weighing 1 mg can generate 1 g of aphid biomass in 23 days and 1 kg in 46 days, under standard culture conditions, as a consequence of their short
FIG. 1 Aphids and other phloem-feeding insects. Aphids, i.e. members of the family Aphididae sensu (Blackman and Eastop, 1984), are one of three families in the superfamily Aphidoidea, within the suborder Sternorrhyncha. All Sternorrhyncha are small (1–10 mm in length) obligate parasites of plants and most (including the Aphididae) feed predominantly or exclusively on plant phloem sap. The Sternorrhyncha are members of the order Hemiptera, which have specialized mouthparts suitable only for liquid diets. The sister group of the Sternorrhyncha comprises the Auchenorrhyncha and Heteroptera, both of which include species that feed on phloem sap or on the cell contents of other plant cells (e.g. mesophyll cells). Other auchenorrhynchans are xylem-feeders and some heteropterans are predacious, using vertebrate blood or other animal fluids. The traditional taxonomy of the Hemiptera treats the Sternorrhyncha and Auchenorrhyncha as sister groups within the suborder Homoptera, but extensive morphological and molecular evidence favours a closer relationship of the Auchenorrhyncha with the Heteroptera than with the Sternorrhyncha (e.g. Goodchild, 1966; Campbell et al., 1995; Von Dohlen and Moran, 1995), as illustrated here.
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larval development time (ca. 1 week) and high reproductive output (ca. 100 live offspring produced over ca. 3-week reproductive period). The parthenogenetic aphids occur naturally in the spring/summer; in temperate conditions, there is a single sexual generation in the autumn, followed by overwintering as coldhardy sexual eggs. This advantage of aphids over other phloem-feeding insects as experimental material has been compounded by the availability of, first, valuable techniques, especially chemically defined diets for nutritional studies (e.g. Dadd, 1985) and electrical recording of feeding behaviour (Tjallingii, 1995) and, second, large datasets, particularly the complete genome sequence of Buchnera in two aphid species, Acyrthosiphon pisum (Shigenobu et al., 2000) and Schizaphis graminum (Tamas et al., 2002). [Genomic data are lacking for aphids, although QTL markers are available for A. pisum (Hawthorne and Via, 2001) and the sequencing of the genome of this aphid species is under consideration.] There is a long tradition of research on the nutritional physiology of aphids (Auclair, 1963; Raven, 1983) but, to the author’s knowledge, Srivastava (1987) is the most recent review. The purpose of this article is to provide an up-to-date overview of the subject, not neglecting the pre-1981 literature that is becoming forgotten through a lack of computerized literature retrieval facilities. However, this article will not consider one important aspect of nutritional physiology, the contribution of physiological approaches to the understanding of aphid ecology, especially plant utilization patterns and life history traits, because this topic has been addressed by Dixon (1998). The opening sections of this article are structured to follow the fate of aphid food, from an overview of plant phloem sap as a diet (Section 2) to aphid acquisition of phloem sap (Section 3) and the processing of ingested sap in the aphid gut (Section 4). Section 5 explores the contribution of symbiotic micro-organisms to aphid nutrition, and the fate of nutrients acquired from both food and symbiotic micro-organisms is reviewed in Section 6. The article concludes on a speculative note by considering likely avenues of future research, including opportunities afforded by recent developments in both analytical methods and molecular biology/genomics (Section 7).
2 2.1
The diet of Aphids: phloem sap PHLOEM SAP TRANSPORT
Phloem sap mediates the bulk flow of photosynthetically fixed organic carbon compounds from ‘source tissues’ (predominantly the leaves) to ‘sink tissues’ (e.g. roots, fruits), at rates of 0.5–3 m per h (Ko¨ckenberger et al., 1997). The transport vessels are highly specialized cylindrical living cells called sieve elements, 20–30 m in diameter, 100–500 m long and aligned end-to-end.
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Adjacent sieve elements are connected by a sieve plate, the pores of which are dilated plasmodesmata (i.e. between-cell connections bounded by plasma membrane). Resistance to sap flow is reduced by large-scale simplification of the cell contents (Van Bel and Knoblauch, 2000). Internal to the cell wall of mature sieve elements are the plasma membrane and a thin parietal cytoplasm containing an endomembrane system, plastids and a few mitochondria all anchored to the plasma membrane, presumably so that they maintain their position. The phloem plastids are not photosynthetically active (they may contribute to sieve plate sealing) and the mitochondria may be non-functional. The nucleus, cytoskeleton, ribosomes and other organelles are absent, and sieve elements are therefore unable to synthesize protein. Each sieve element is metabolically and genetically dependent on its sister cell, called the companion cell, to which it is connected by specialized cytoplasmic pores, known as the pore-plasmodesmal units, through which small proteins can pass (the molecular exclusion limit is 10–40 kDa) (Kempers and van Bel, 1997). Apart from this cytoplasmic link to companion cells, transport sieve elements are almost completely isolated, i.e., they approximate to leakfree pipes. The osmotic pressure in the sieve elements is 0.6–3 MPa, and the flow of phloem sap is widely accepted to be under positive pressure and driven by a greater osmotic pressure in the source tissues than the sink tissues. The implications are, first, that the concentration of phloem sap solutes in one sieve tube declines progressively with distance from the source tissue; and, second, that the rate and direction of phloem sap movement varies both with position in the plant and over time, according to the relative strength of sources and sinks. As a consequence, the nutrients available to insects feeding at different locations along one sieve tube or from different sieve tubes of one plant are not identical. Generally, the photoassimilates are loaded into the sieve element/companion cell complex in the minor veins and at the vein endings of the source tissue. However, sieve elements are absent from the tips of the veins. For example, in leaves of the potato, sieve elements extend to the seventh-order veins of the abaxial surface and sixth order veins of the adaxial surface (McCauley and Evert, 1989). According to the mass flow hypothesis of phloem translocation as originally envisaged by Mu¨nch (see Fisher, 2000), all plant cells from the site of photosynthetic fixation in the source tissues to the sieve elements and thence to the sink tissues are hydraulically connected. Many plants, including an estimated 80% of all woody species, approximate to this condition. They are called ‘symplasmic loaders’, and the photoassimilates are transferred to the sieve elements in minor veins of the source tissues via numerous plasmodesmal connections between the cells (Fig. 2). Many plants, however, have very few plasmodesmal connections between the sieve-element/companion cell complex and adjacent cells in the source tissue, and the solutes are loaded into the sieve element by uptake from the apoplast (extracellular space) across the plasma
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FIG. 2 Long-distance transport of photoassimilates via the sieve elements of vascular plants. Sieve elements are in symplasmic connection with each other, companion cells and the sink tissue via plasmodesmata. Phloem loading at the source tissue is mediated by two alternative processes: (a) symplasmic loading: sucrose is transported symplasmically from the site of synthesis to the sieve element. In the companion cell, some of the sucrose is transformed to raffinose and other sugars of the raffinose series, which are loaded into the sieve element via the plasmodesmatal pore, but are too large for the transport via standard plasmodesmata in the reverse direction. The proposed accumulation of raffinose series oligosaccharides in the sieve element is known as polymer trapping (see Section 2.2.1 for details); and (b) apoplastic loading: sucrose is released into the apoplast in minor veins and taken up by the H þ /sucrase transporter (shown as a closed circle) located on the cell membrane of the companion cell (e.g. Arabidopsis thaliana) or sieve element (e.g. potato).
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membrane. Apoplastic loading is evolutionarily more derived than symplasmic loading (Turgeon et al., 2001), and is restricted largely to herbaceous angiosperms, especially species in cold and dry climatic zones (Van Bel and Gamalei, 1992); temperate crops are generally apoplastic loaders. Some taxa display structural, and probably functional, variation in the sieve elements. Notably, the Graminae have both thick-walled sieve elements, which function principally to retrieve minerals and solutes from xylem, and thinwalled sieve elements, which mediate long-distance transport (Fritz et al., 1983). The aphid Sitobion yakini preferentially feeds from the latter (Matsiliza and Botha, 2002). 2.2
THE COMPOSITION OF PHLOEM SAP
Phloem sap is sampled by three methods: exudation from shallow incisions into the stem, exudation from cut surfaces into EDTA solution (which inhibits sieve element sealing) and from the severed stylets of aphids (Kennedy and Mittler, 1953; King and Zeevart, 1974; Peel, 1975). The last method is strongly preferred because the sap composition is not contaminated by the contents of other cell types or confounded by the plant response to wounding. Its one major drawback is that, by definition, it cannot control for possible aphidinduced changes in phloem sap content (see Section 2.3), but this is more of a problem for plant physiologists than for insect physiologists interested in phloem sap feeders. The limitations of the EDTA exudation technique are well defined (Weibull et al., 1990). The EDTA exudates provide an excellent measure of the amino acid composition and amino acid:sugar ratio, but not of either total amino acid content or sugar composition (because phloem sugars are hydrolysed by contaminating enzymes in the exudates). 2.2.1
Carbohydrates
The dominant compounds in phloem sap are carbohydrates, especially sugars. Sucrose is a major transport sugar in most plants, probably linked to its chemical stability (it is a non-reducing sugar) and low viscosity in concentrated solutions (Lucas and Madore, 1988). It accounts for 95% or more of the phloem sugars in species with apoplastic phloem loading (see Section 2.1), reaching concentrations of 0.5 M–1.5 M. This is an order of magnitude higher than in the cytoplasm of other plant cells (Winter et al., 1992), and is a consequence of the active transport of sucrose into the sieve element– companion cell complex (Ward et al., 1998) (Fig. 2). In symplasmic loading plants with sucrose as the dominant phloem sugar (e.g. Populus, Salix), the sucrose concentration is not elevated in the sieve elements relative to mesophyll cells (Turgeon and Medville, 1998). Over the diurnal cycle, the phloem sucrose
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concentration generally varies by less than two-fold because the phloem sucrose is derived from both recently assimilated carbon and, especially at night, from starch accumulated during the day in the source tissues (Winter et al., 1992; Genger and Servaites, 1994; Kehr et al., 1998). Sucrose is not the sole phloem sugar in many plants. In particular, some taxa, notably members of the Lamiaceae and Cucurbitaceae, also transport sugars of the raffinose family, including raffinose, stachyose and higher order oligosaccharides, synthesized by the stepwise transfer of one, two or more galactose units, respectively, to the glucose moiety of sucrose. Although plants with phloem sugars of the raffinose series are generally symplasmic loaders, the raffinose sugars are loaded against a concentration gradient and, therefore, can attain high concentrations in the phloem sap. This apparent paradox has been accounted for by the polymer trapping model of Turgeon (1991), as illustrated in Fig. 2a. Polyols (sugar alcohols) are also quantitatively important phloem compounds in several families of dicots: sorbitol in Rosaceae and Plantaginaceae, galactitol ( ¼ dulcitol) in the Celestraceae and Scrophulariaceae, and mannitol in many species including members of Oleaceae and Apiaceae (Noiraud et al., 2001). As examples, the principal carbohydrates in the phloem sap of apple trees are sorbitol (65–75%) and sucrose (25–35%) (Klages et al., 2001); the concentration of mannitol in celery phloem sap is 150–300 mM, accounting for up to 60% of the total phloem carbohydrate (Hu et al., 1997); and the dominant phloem carbohydrates in Primula are an unusual seven-carbon polyol, volemitol (D-glycero-D-manno-heptitol) and sucrose (Ha¨flinger et al., 1999). The contribution of polyols to phloem carbohydrates may be particularly important during the night (Klages et al., 2001) and their absolute concentrations are elevated under drought stress and at elevated temperature in some plant species (Noiraud et al., 2001). 2.2.2
Lipids and sterols
It is widely assumed that lipids and sterols are not quantitatively important components of phloem sap. However, this is based on rather little experimental data; the comment of Ziegler (1975) ‘there are very few analyses, none of which are systematic, of the lipid content of the sieve tubes’ provides a fair description of the current literature on phloem lipids. Exceptionally, Mady et al. (2002) detected free fatty acids and, at lower concentrations, triacylglycerols, phospholipids and steryl and wax esters in EDTA exudates from oil-seed rape (Brassica napus). The dominant fatty acid was palmitic acid (16:0), appreciable amounts of lauric acid (12:0), myristic acid (14:0) and pentadecanoic acid (15:0) were also present, and the ratio of saturated:unsaturated fatty acids was about five-fold higher than in the shoot cytosol fractions. Absolute concentrations cannot be obtained by this method. Although the authors present evidence that the exudates were not
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contaminated by non-phloem material and that unsaturated fatty acids were not oxidized to saturated forms during exudation, these results should be verified with stylet exudate data and a range of plant taxa. Plant sterols are chemically distinctive in that they are alkylated with either C1 or C2 substituents at C-24 of the parent molecule. Common plant sterols, including sitosterol (C29), stigmasterol (C29) and campesterol (C28), are phloem-mobile, as revealed by their recovery in the honeydew of phloemfeeding insects reared on plants (Forrest and Knight, 1972; Campbell and Nes, 1983). The impact of environmental conditions and plant developmental age on the concentration and composition of lipids and sterols in phloem sap is not wellknown. O. Biddulph and R. Cory are reported in Ziegler (1975) to have found detectable phloem sterols in young, but not old, leaves of soy bean Glycine max; and Mady et al. (2002) describe substantial increased fatty acid levels in EDTA exudates of B. napus after exposure to UV-irradiation. 2.2.3
Nitrogenous nutrients
The principal nitrogenous compounds in the phloem sap of most plants are free amino acids, with a total concentration between 50 and 800 mM; the concentration of inorganic nitrogenous compounds, ammonium and nitrate, are very low and undetectable, respectively. Many, often all, of the 20 amino acids which contribute to protein are detectable (i.e. at concentration >1 nM), and the dominant amino acids are usually one-to-all of aspartate, glutamate, asparagine and glutamine. The concentration of ‘essential’ amino acids, i.e. amino acids which animals cannot synthesize de novo, is low, accounting for 10–30% of the total protein amino acids. (The essential amino acids are: histidine, isoleucine, leucine, lysine, methionine, phenylalanine, threonine, tryptophan and valine.) Within these bounds, the total concentration and composition of phloem amino acids are very variable. It is commonplace to obtain three-to-five-fold variation in amino acid concentration among replicate samples of exudates from severed stylets using plants raised under tightly controlled conditions (Girousse et al., 1996; Telang et al., 1999) and variation over the diurnal cycle is also reported (Winter et al., 1992; Corbesier et al., 2001). The composition varies considerably with plant developmental age (Corbesier et al., 2001; Karley et al., 2002), environmental conditions, especially drought stress (Girousse et al., 1996), season (Douglas, 1993; Sandstro¨m, 2000) and between plant species (Ziegler, 1975; Wilkinson and Douglas, 2003). In barley, which has been studied intensively, the total concentration of amino acids in the cytoplasm of leaf mesophyll cells and phloem sap are equivalent, suggesting that amino acid transfer to the sieve elements is passive, even though phloem loading is apoplastic in this species (Winter et al., 1992). However, the composition of amino acids in the two locations is not
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FIG. 3 A comparison of two indices of leaf nitrogen content in potato plants: total leaf carbon-to-nitrogen ratio (C:N, closed circles) and molar ratio of sucrose to amino acids (sucrose:amino acid, open circles) in leaf phloem exudates. (Reproduced from Fig. 1b of Karley et al. (2002).)
equivalent, indicative of some selectivity in amino acid loading. Synthesis of glutamine from glutamate via glutamine synthetase activity in the companion cell/sieve element complex may additionally contribute to the high glutamine levels in the phloem sap of certain species (Pereira et al., 1992). Linked to the active accumulation of sucrose but not amino acids in the sieve elements of apoplastic loading plant species, the sucrose:amino acid ratio (mol/mol) in phloem sap, at 3:1–20:1 is considerably higher than in the cytoplasm of other plant cells, where the ratio is ca. 1:1. As a result, there is little justification for the widespread use of total plant tissue C:N as an index of the nutritional value of plants for phloem sap-feeding insects. For example, the total leaf C:N and phloem sucrose:amino acid ratio of the potato Solanum tuberosum are not well correlated and vary in different ways with plant developmental age (Fig. 3). Non-protein amino acids, peptides and various other nitrogenous compounds have been reported in plant phloem sap. In particular, the sulphur amino acids methionine and cysteine are transported principally as the nonprotein amino acid S-methylmethionine and the tripeptide glutathione (glutamate–cysteine–glycine), respectively, which together can account for 3.5% of the total phloem amino acids and represent the principal route for phloem transport of sulphur (Rennenberg, 1982; Bourgis et al., 1999). Other nitrogenous compounds, including polyamines and ureides, have been identified in the phloem sap of various plants (Ziegler, 1975), and some limited data are available on the variation in their concentration with environmental conditions and plant developmental age (Antognoni et al., 1998). The phloem proteins encountered by phloem-feeding insects are known as the sieve tube exudate proteins (STEPs), which comprise at least
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150–200 different proteins, mostly of molecular weight 10–40 kDa and at a very low total concentration, 0.1–2 g l 1. Exceptionally, the Cucurbitaceae have protein-rich phloem sap at 10–40 g l 1. Most STEPs are synthesized in companion cells and translocated into the sieve elements, and they may be cycled back to the companion cells for degradation or transported to sink tissues (Thompson and Schulz, 1999). Hayashi et al. (2000) suggest that phloem proteins have roles in: (a) intermediary metabolism, e.g. all enzymes in glycolysis are detectable in phloem sap, suggesting that sieve elements may contribute to ATP production; (b) redox regulation, as mediated by thioredoxin h, glutaredoxin and glutathione reductase, all of which have been detected in phloem sap; and (c) defence against insect and microbial attack (see Section 2.2.5). In cucurbits (which have high phloem protein concentrations), they may also contribute to the long-distance transport of nitrogen. The sieve elements of many species also bear structural proteins, known as P-proteins, in the parietal cytoplasm. These proteins do not generally occur in the sap stream, although some, which form crystalline or amorphous accumulations (called P-protein bodies) in the cytoplasm, may be in equilibrium with the soluble monomeric form in the STEP fraction. The P-proteins have been implicated in the wound response of sieve elements, especially in legumes. There is some evidence to support the notion that the sieve elements respond to loss of turgor pressure by increased cytoplasmic concentration of Ca2 þ ions (released from the endoplasmic reticulum or taken up from the apoplast), resulting in the dispersal of the cytoplasmic P-proteins and their plugging of the pore plates (Van Bel et al., 2002). This response may be complemented by (and in taxa apparently lacking P-proteins, replaced by) the deposition of callose at the pore plates, but there is some evidence that callose deposition may be an artefact of preparation methods for microscopical analysis (Radford and Overall, 2001). 2.2.4
Inorganic substances
The total concentration of inorganic substances in phloem sap is 1–5 g l 1, and the dominant components are potassium, K þ , at 50–150 mM, and phosphate, at 10–50 mM. K þ ions are crucial to the phloem transport of photoassimilates, as indicated by the depressed solute loading and phloem flow, and linked sucrose accumulation in source leaves of K þ -deficient plants (Kochian, 2000). Phosphate and other anions, including bicarbonate and organic acids (e.g. malate), maintain the charge balance and play a large role in determining the pH of the phloem sap, at 7.3–8.0 units (Ziegler, 1975; Marschner et al., 1996; Fisher, 2000). The other principal phloem-mobile inorganic ions are Mg2 þ , at ca. 5 mM, and Cl , at ca. 10 mM (but see below). The concentrations of inorganic nitrogen (as nitrate and ammonium) and sulphur (as sulphate) are low, often undetectable, in phloem sap, and the
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elements nitrogen and sulphur are transported principally as organic compounds (see Section 2.2.2). The calcium concentration in phloem sap is also low, at 10 M–3 mM, of which 10–30 M is free and the remainder is chelated (Fisher, 2000; Volk and Franceschi, 2000). Phloem sap also contains detectable amounts of the micronutrients, including iron, manganese, zinc, copper and molybdenum. Boron is generally considered to be phloemimmobile, but its phloem mobility is reported to be enhanced by the presence of polyols in the phloem sap (Noiraud et al., 2001). The Na þ :K þ ratio of phloem sap is generally low, at 1:5–1:10 (Fisher, 2000). Plant biologists dispute whether sodium is an essential nutrient for plants, and high sodium levels are undoubtedly toxic to many plant species. Na þ is, however, phloem-mobile and the concentration of Na þ in phloem sap is elevated when plants are grown under saline conditions. For example, when Lohaus et al. (2000) watered maize plants with 0.1 M NaCl, the phloem Na þ concentration was >10 mM, an order of magnitude higher than that in plants watered with distilled water; the high salt treatment increased the phloem Cl concentration ca. three-fold, from 8 to 30 mM, but had little effect on the concentration of sucrose, amino acids and other inorganic ions. 2.2.5
Defences against insects
Plant chemical defences against herbivory principally comprise toxins and compounds which interfere with digestion (e.g. tannins). They are located in the apoplast and cell vacuole, where they have no deleterious effects on plant metabolism, and are excluded from sieve elements, which are a cytoplasmic compartment bounded by the plasma membrane. As a consequence, phloem sap poses fewer toxicological hazards for animals than most other plant tissues. However, the phloem sap of some plant species is not entirely without chemical protection. Analysis of both phloem sap and the tissues and honeydew of feeding aphids offer unambiguous evidence for phloem-mediated transport of various secondary compounds. These include: glucosinolates, found mainly in crucifers and other Capparales (Merritt, 1996; Brudenell et al., 1999; Chen et al., 2001); cardenolides, particularly in the Asclepiadaceae (Botha et al., 1977); alkaloids, e.g. ricinine (Waller and Skursky, 1972), quinolizidines (Wink and Witte, 1991) and pyrrolizidines (Hartmann, 1999), in various plant groups; and the glucoside of hydroxamic acid in wheat (Givovich et al., 1994). The concentration of secondary compounds in the phloem sap varies with developmental age of the plant and, at least for the quinolizidine alkaloids, with time of day (Wink and Witte, 1991). The sieve elements of all plants are well-defended against mechanical damage. When a sieve element is damaged, the sieve pores are plugged, preventing any further phloem flow through the affected sieve tube. The plugging material may include phloem P-proteins, sieve element plastids (which
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‘explode’ on wounding) and callose, but the relative importance of these different components is disputed and may vary between plant species. Calcium may be a crucial signal in the sieve element wounding response, when the free Ca2 þ concentration is probably transiently elevated (by release from stores in the endoplasmic reticulum of the sieve elements or uptake from the apoplast), resulting in sealing of plasmodesmata and sieve plates (Van Bel et al., 2002). When aphids imbibe phloem sap from susceptible plants, the structural organization of the sieve elements is not usually altered detectably, at either the light- or electron-microscopical levels, suggesting that feeding may depend on circumventing or suppressing the sieve plate sealing response. Consistent with this interpretation, sieve element sealing has been implicated as a possible resistance factor in certain plant/aphid combinations (Van Helden and Tjallingii, 1993; Caillaud and Niemeyer, 1996). 2.3
APHID-INDUCED CHANGES TO PHLOEM SAP
Insects feeding on phloem sap are hydraulically equivalent to plant sinks, such as fruits or roots. There is, however, evidence that the relationship between an aphid and a susceptible plant is more complex than the relationship between the source tissues and sink tissues of a plant. Various studies indicate, first, that aphid feeding induces specific plant responses and, second, that these responses can include changes in phloem sap that are advantageous to the insect. Aphid feeding induces changes in the patterns of plant transcription and protein synthesis that overlap with inducible plant responses to microbial pathogens and are distinct from plant responses to mechanical wounding as evoked by chewing insects (Walling, 2000; Gatehouse, 2002). For example, Arabidopsis thaliana plants infested with the green peach aphid Myzus persicae display a strong response in the salicylic acid-pathway, including an order of magnitude increase in the levels of the pathogenesis-related gene PR-1 and the -1,3,-glucanase gene BGL2 (Moran and Thompson, 2001). These responses are traditionally considered as defensive. However, they are largely localized to the plant apoplast, not the contents of the sieve elements, and they do not translate into resistance to the aphid. Indeed, some aphid-induced changes in the plant tissues may promote phloem suitability for the insect. For example, Moran and Thompson (2001) suggest, first, that the elevated activity of glucanase (the product of BGL2) near phloem sieve elements may counteract callose deposition, a putative sieve element wound response (see Section 2.2.5) and, second, that the increased transcription of the gene STP4, coding a sugar-H þ transporter, may promote the insect-sink. The possibility that aphids may modify the nutrient composition of plant phloem sap has been discussed extensively in the literature (Klingauf, 1987; Dixon, 1998). If an aphid induced changes in the sieve element from which it
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was feeding, one would anticipate systematic differences between the composition of phloem exudates from severed aphid stylets and from EDTA exudates, which sample all the sieve elements of the plant tissue being tested. No such differences have been identified (Weibull et al., 1990), suggesting that, contrary to suggestions in the literature, aphids generally do not manipulate phloem sap composition at the level of the single sieve element. Exceptionally, aphids which induce chlorotic lesions or galls on their food plants may modify plant sap composition. Considering aphid-induced chlorosis first, Sandstro¨m et al. (2000) compared the amino acid profiles of exudates from severed stylets of three aphid species feeding on wheat: Rhopalosiphum padi, which causes no visible macroscopic changes to the plant, and S. graminum and Diuraphis noxia, which cause leaf necrosis and chlorosis. Both the total concentration and the composition of phloem amino acids varied significantly among the three species, with elevated concentrations of amino acids, especially essential amino acids, in the exudates from S. graminum and D. noxia stylets, relative to R. padi (Fig. 4). Further experiments revealed that the amino acid composition of EDTA exudates from leaves uninfested with aphids were closely similar to those of leaves bearing R. padi and different from those with S. graminum (Sandstro¨m et al., 2000), and that D. noxia reared on a resistant cultivar of wheat induces neither lesions nor a change in phloem composition (Telang et al., 1999). Several conclusions can be drawn: that feeding by S. graminum and D. noxia enhances the nutritional quality of wheat phloem sap; that aphidinduced changes in phloem sap are systemic, at least to the level of the infested
FIG. 4 Amino acid content of phloem sap exudates from severed stylets of aphids feeding from wheat seedlings (closed bar, essential amino acids; open bar, non-essential amino acids): the mean % essential amino acids in exudates from Rhopalosiphum padi is 29%, Diuraphis noxia is 33% and Schizaphis graminum is 42% (essential amino acids are defined as the nine amino acids that cannot be synthesized de novo by animals). (Figure drawn from data in Sandstro¨m et al. (2000).)
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leaf; and that these changes are correlated with aphid-mediated induction of lesions. The impact of lesion-inducing aphids on plant phloem sap bears parallels to the indirect interactions between plant pathogenic fungi and aphids. In particular, improved leaf nutritional quality, arising from leaf chlorosis and premature leaf senescence induced by fungal infection, has been shown to result in improved performance of the alder aphid Pterocallis alni on leaves of alder Alnus pubescens (Gange, 1995) and of the birch aphid Euceraphis betulae on leaves of birch Betula pendula (Johnson et al., 2003). The phloem sap of senescencing leaves is enriched in amino acids derived from hydrolysis of leaf proteins prior to abscission. However, aphids are believed to induce chlorotic lesions by salivary toxins or enzymes (Miles, 1999) and the extent to which the processes underlying the resultant induced shifts in phloem composition match those underlying fungal-promoted or ‘natural’ senescence remains to be established. Various aphids induce plant galls. It is widely accepted that the phloem sap is nutritionally superior in galled tissue than ungalled tissue (Forrest, 1987; Dixon, 1998), although this difference has not been quantified directly. The underlying plant physiological processes are also obscure and probably different from chlorosis-linked shifts in plant sap composition because galls are persistent structures and not subject to premature senescence.
3 3.1
Acquisition of phloem sap by aphids INGESTION OF PHLOEM SAP
As in other hemipteran insects, the mandibles and maxillae of aphids are transformed into needle-like stylets, the mandibular stylets surrounding the inner maxillary stylets. Two canals, a salivary canal (0.2–0.4 m diameter) and larger food canal (1–2 m diameter), lie between the tightly apposed maxillary stylets; and each mandibular stylet contains one canal bearing two nerve axons (Fig. 5a). When the insect is not feeding, the stylets are enclosed within the labium (also known as the rostrum or proboscis). Aphids lack maxillary and labial palps, and chemoreception is mediated by sensilla in two locations: at the tip of the labium and in the epipharyngeal gustatory organ (Fig. 5b). Plant phloem sap in the sieve tubes is under positive pressure (see Section 2.1), sufficient for it to be forced up the food canal of the stylets into the pharyngeal duct of the aphid. The empirical evidence for passive ingestion is that phloem sap continues to exude from severed stylets at a rate comparable to the rate of honeydew production by the feeding insect (Auclair, 1963). However, the insect does have some control over the rate of phloem sap ingestion (Ponsen 1987). The food passes into the alimentary tract only if the pharyngeal valve, which lies just distal to the epipharyngeal gustatory organ, is
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FIG. 5 The mouthparts and foregut of an aphid. (a) Diagrammatic transverse section through the stylets; (b) longitudinal section through head to show cibarial pump and head musculature (shaded), with the detail of the pharyngeal valve (abbreviated to ‘valve’ on figure) displayed as the inset: the salivary canal of the stylets and the salivary glands are not shown for clarity. (Redrawn from Fig. 2.3 and Fig. 2.7 of Dixon (1998).)
open, a condition that requires the contraction of dilator muscles in the aphid head. An aphid can, presumably, halt ingestion very rapidly in response to noxious or deterrent compounds detected by the epipharyngeal gustatory organ. Distal to the pharyngeal valve is the cibarial pump (also known as the pharyngeal pump), which, as the name suggests, has the opposite capability: it can ‘suck’. The dorsal wall of the cibarial pump is flexible and, when dilator muscles attached to the dorsal wall contract, the volume of the pump is
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increased, drawing fluid in. When the muscles relax, the liquid in the pump is either forced distally down the alimentary tract (when the pharyngeal valve is closed) or ejected back down the stylets (when the pharyngeal valve is open). An indication of the capability of the cibarial pump comes from the capacity of aphids to suck liquid from artificial diets at atmospheric pressure for days-to-weeks and from plant xylem vessels at pressure of 0.02–0.2 MPa for at least several hours (Spiller et al., 1985). For most animals, the amount of food ingested is determined by the duration of feeding bouts and the ingestion rate during each feeding bout. On susceptible plants, aphids often feed continuously from single sieve elements for extended periods. The evidence comes principally from two complementary sources. The first is ‘honeydew clock’ experiments, where each honeydew droplet is collected on paper on a slowly moving rotating table placed below a feeding insect; aphids produce honeydew droplets of uniform size at regular intervals of once per 15–40 min, varying with aphid and plant species. The second is electrical monitoring of aphid feeding, in which an aphid is part of an electric circuit and the position of its stylet tips and stylet activity can be determined from the associated changes in resistance, recorded as changes in potential on an electrical penetration graph (EPG) (Tjallingii, 1995). EPG traces of aphids feeding from susceptible plants generally include many uninterrupted hours of the wave pattern associated with ingestion of phloem sap from sieve elements. These data have led to the consensus view that aphids do not have well-defined feeding bouts, dictated by temporal changes in appetite and satiation (Tjallingii, 1995); but for a different perspective, see Montllor (1991). Aphid feeding rates increase with body size, both over larval development (Febvay et al., 1999) and among aphid species (Dixon, 1998). Generally, aphids feed faster on the phloem sap of susceptible plants than on artificial diets. Peel (1975) estimated that severed stylets may exude ca. 1 l of phloem sap h 1, which is equivalent to emptying 235 sieve elements per minute, and Pollard (1973) calculated that M. persicae ingests 10% of its body weight h 1. Aphid feeding rates also vary with the nutrient content of the ingested food. Early studies with diet-reared aphids demonstrated that sustained feeding depended on the presence of sucrose at concentrations >0.1 M in the diet and was promoted by at least one amino acid, methionine. Sucrose and methionine are, therefore, widely regarded as phagostimulants. In addition, aphids exhibit compensatory feeding responses, i.e. elevated feeding rates on dilute diets, resulting in increased nutrient uptake. Compensatory feeding on diets containing low sucrose concentrations has been described for both M. persicae and A. pisum (Mittler and Meikle, 1991; Abisgold et al., 1994). In A. pisum, increased feeding rates only partly compensates for low dietary concentrations, such that the total amount of sucrose ingested declines with decreasing sucrose concentration (Fig. 6). Aphids also display compensatory feeding response to dietary amino acid levels at concentrations in the range 75–250 mM, but not at higher concentrations (Prosser et al., 1992; Abisgold
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FIG. 6 Feeding response of pea aphids Acyrthosiphon pisum to variation in concentration of dietary sucrose. Eight replicate final-instar apterous larvae were caged individually for 48 h to diets containing 0.2–1.0 M sucrose. (a) Volume of diet ingested. (b) Amount of sucrose ingested (circle) and recovered from aphid tissues (square) and honeydew (triangle). (Previously unpublished data of W.A. Smith and A.E. Douglas; methodology as in Ashford et al. (2000).)
et al., 1994), and their feeding responses to other dietary constituents, notably inorganic ions, have not been investigated. 3.2
STYLET PENETRATION TO THE SIEVE ELEMENTS
Stylet penetration to the sieve elements is predominantly intercellular. The stylets pass directly through the plant cell walls, a route that is known as the intramural pathway, and not along the middle lamella between the walls of adjacent cells. From the time when the aphid stylets penetrate the plant surface, it takes between 20 min and more than an hour for them to reach a sieve element and the aphid to initiate feeding; this period of stylet penetration through the plant tissues is commonly called ‘the pathway phase’. Complementary microscopical analysis and EPG studies indicate that the stylets puncture plant cells briefly (usually for <10 s) during the pathway phase (Tjallingii, 1995). The significance of these penetrations into the cytoplasm of plant cells is uncertain. Perhaps, they reflect the difficulties in maintaining the
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intramural pathway (i.e. the aphid stylets are ‘clumsy’), and perhaps they provide the opportunity for aphids to sample cell contents, as a check on the suitability of the plant or in search for a sieve element. Consistent with the latter perspective, plant cytoplasmic contents are imbibed during the brief puncture, at a rate of ca. 0.1 pl s 1 (Martin et al., 1997). The punctures do not cause damage to the plant cell evident at the microscopical level, but they may trigger plant changes, for example in gene expression or at the biochemical level (see Section 2.3). The factors determining the direction of stylet progression through the plant tissue are not understood. Microscopical analyses indicate that the stylet pathway is irregular, often with multiple changes in direction, and does not take the shortest route to the plant vascular tissue (Tjallingii and Esch, 1993). These data suggest that the plant sieve elements are located by trial-and-error. Furthermore, an aphid does not necessarily initiate sustained feeding from the first sieve element penetrated. This raises two linked questions: do sieve elements in one plant vary in their suitability for aphids, and what cues does an aphid use to initiate sustained feeding from a sieve element? Perhaps, stylet movement is arrested principally by the high osmotic pressure and sugar concentration in sieve elements. Other phloem constituents may also influence sieve element suitability. For example the possible significance of phloem amino acid levels is suggested by the finding of Ponder et al. (2000) that the stylets of R. padi feeding on barley penetrate to the sieve elements more times for every incidence of sustained feeding on nitrogen-deficient plants, with 110 14 mM phloem amino acids, than on nitrogen-sufficient plants, with 180 23 mM phloem amino acids. Aphid saliva is central to the progression of aphid stylets through plant tissues to the sieve elements, and initiation of sustained feeding, and this issue is considered in the next section. 3.3
APHID SALIVA
The production and composition of saliva has been studied over many years in aphids, and Miles (1999) provides a comprehensive review. Aphid saliva is produced in paired salivary glands and secreted via the salivary canal. (As considered in Section 3.1 and Fig. 5a, the salivary canal is one of two canals between the tightly apposed maxillary stylets; the other canal is the food canal, along which the liquid food is passed in the opposite direction.) The food canal, but not the salivary canal, extends to the distal tips of the maxillary stylets and, therefore, saliva is predicted to exit the stylets into the plant tissues only when food is not being ingested (Prado and Tjallingii, 1994). Two types of aphid saliva are recognized: the gelling saliva produced as the stylets penetrate through plant tissues but not once the stylet tip accesses a sieve element, and the watery saliva produced during both penetration and the sieve element phase. The gelling saliva is a lipoprotein of unknown
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biochemical composition, which solidifies on extrusion from the stylets, principally because sulphydryl groups are oxidized to form disulphide bonds that promote hydrogen bonding. The watery saliva, which has pH 8–9, contains free amino acids, various proteins, and reducing activity of unknown chemical basis. The gelling saliva is crucial to stylet penetration through the plant tissues. It forms a continuous tube, or sheath, around the stylets as they extend through the plant tissues, and it may both support the stylets mechanically and protect them from plant defence responses. When the aphid retracts its stylets from a plant, the stylet sheath remains undisturbed in the plant tissue, leaving a track of the movement of stylets through the plant tissue. The watery saliva contains various enzymes, including polyphenol oxidase ( ¼ catechol oxidase) which may detoxify phloem-mobile allelochemicals, and pectinesterase, polygalacturonase and cellulase which have been suggested to promote stylet penetration through plant cell walls and to degrade plant oligosaccharides produced by aphid wounding to inert monomers (Miles, 1999). It is, however, widely accepted that aphid stylet penetration has a predominantly mechanical basis, with little or no contribution from enzymatic degradation of the plant cell wall material. The watery saliva has also been suggested to ‘condition’ the sieve elements, perhaps linked to preventing plant processes that halt phloem flow in damaged sieve elements (e.g. callose formation, P-protein precipitation). The basis of this putative conditioning is chemically undefined and it is believed to occur in the first 30 seconds-to-few minutes after sieve element puncture by the stylets, when the aphids are salivating but not ingesting phloem sap (represented by E1 waveform in EPG analyses). Consistent with a possible role of salivation in phloem conditioning, some aphids feeding from plants on which they perform relatively poorly salivate into the sieve element for extended periods (>5 min) before initiating sustained feeding, and also make multiple transitions between phloem ingestion and salivation during the subsequent hours when the stylets are tapped into the sieve element (Van Helden and Tjallingii, 1993). Aphid-mediated ‘conditioning’ of plants is considered further in Section 7.1.1. There are several early reports of -glucosidase and -fructosidase activities in aphid saliva (reviewed in Srivastava, 1987). In principle, such enzymes could initiate digestion of phloem sugars before the sap reaches the gut. However, the purity of the saliva extracts and specificity of the enzymatic assays in these studies are uncertain (Miles, 1999), and further research is required to validate the results and demonstrate their quantitative importance.
4
Processing of food in the aphid alimentary tract
Feeding aphids ingest a continuous stream of phloem sap, a liquid food of high osmotic pressure comprising nutrients of low molecular weight and of
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unbalanced composition (e.g. high sucrose:amino acid ratio and high ratios of K þ :Na þ and Mg2 þ :Ca2 þ ). This diet generates three predictions about the function of the aphid alimentary tract: (1) It mediates the efficient assimilation of limiting organic nutrients and inorganic ions, such as amino acids and Ca2 þ ions; (2) it has a restricted digestive capacity, largely related to the hydrolysis of di- and oligo-saccharides to monosaccharide sugars; and (3) it contributes to the maintenance of osmotic balance and regulation of the water content of the insect. Our understanding of all three issues is only fragmentary, however, because physiological analysis of the aphid alimentary tract is technically challenging. The gut is small and delicate, largely precluding physiological studies of isolated gut preparations, and limiting the material available for purification and biochemical analysis of gut enzymes. Most research on gut function has, to date, been conducted on intact insects, which are amenable to two approaches: quantification of assimilation efficiency and analysis of chemical transformations in ingested food in the gut. These methodologies are considered further in Section 4.1, followed by a review of our current understanding of the structural organization of the aphid alimentary tract (Section 4.2), the processing of sugars and nitrogenous compounds (Section 4.3 and Section 4.4) and the fate of ingested allelochemicals (Section 4.5). The gut-mediated processing of ingested sugars is linked to osmoregulation, and these two topics are therefore addressed together (see Section 4.3). The translocation of inorganic ions across the gut epithelium and the significance of the gut, especially the rectum, in ionic regulation have not been studied in aphids. 4.1 4.1.1
APPROACHES TO STUDY FOOD PROCESSING
Assimilation efficiency
The assimilation efficiency of a nutrient refers to the proportion of an ingested nutrient that is assimilated across the gut wall into the insect tissues. The amount ingested per unit time is determined from the product of feeding rate and dietary concentration of the nutrient, and the amount assimilated is the difference between the amount ingested and recovered in the egesta (i.e. in the honeydew released from the aphid). The assimilation efficiency of aphids reared on chemically defined diets can be determined readily because the dietary concentration is known and feeding rate can be obtained from the quantitative recovery in the honeydew of a non-metabolizable compound included in the diet at a defined concentration (radioactively labelled inulin is widely used as the non-metabolizable compound (Wright et al., 1985)). The values of assimilation efficiency obtained for diet-reared aphids should be extrapolated to aphids on their natural food of plant phloem sap with the greatest caution. Aphid feeding rate is generally lower on diets than on plants,
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and it has been suggested that the assimilation of nutrients from phloem sap may be limited by its short transit time in the gut (Dixon, 1998). This issue has not, to my knowledge, been investigated experimentally. It is technically demanding to obtain accurate values of aphid feeding rates on plants and nutrient concentrations of phloem sap in the sieve tubes from which an aphid is feeding, and few studies have included estimates of the assimilation efficiency of aphids feeding from plants. More generally, the limitations of assimilation efficiency as an index of the food processing capability of an animal have been identified (Raubenheimer and Simpson, 1994). In particular, because this index is a proportion, it can provide an incomplete and potentially misleading measure of the amount of a nutrient available to insect tissues. Information on the absolute rates of nutrient assimilation, as well as the assimilation efficiency, is increasingly being provided in nutritional physiology studies of insects, including aphids. 4.1.2
Chemical transformations of ingested food in the gut
Aphids reared on chemically defined diets have also been a vital tool in the analysis of digestion and other gut-mediated chemical changes to ingested compounds. A productive approach is to compare the composition of honeydew produced by aphids feeding on diets containing the compound(s) of interest at different concentrations. The dietary compound(s) may be labelled, for example with radioactivity, to define more precisely the details of the chemical changes. The chief caveat with this whole-organism approach is that the site of chemical transformation is not defined. Aphid honeydew is not simply the residue of the ingesta after digestion and assimilation in the gut, but also contains waste products of aphid metabolism voided via the gut. For example, aphids excrete ammonia in the honeydew and, when reared on diets with individual amino acids deletions, their honeydew commonly contains the amino acid absent from the diet, varying among both amino acids and aphid species (Douglas and Prosser, 1992). One approach to investigate whether the chemical transformation is gut-mediated is to conduct parallel experiments with dissected guts in insect saline (Douglas, 1990) or with crude homogenates of guts (Ashford et al., 2000). Such experiments, however, need careful interpretation because the isolation procedure may destroy the structural organization of the alimentary tract or result in the degradation of key enzymes. 4.2
STRUCTURAL ORGANIZATION OF THE ALIMENTARY TRACT
The alimentary tract of aphids is simple anatomically (Fig. 7). Formally, the ‘mouth’ of the aphid is at the junction of the cibarial pump and the oesophagus, which is a thin-walled tube opening, via the oesophageal valve,
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FIG. 7 The alimentary tract of the green peach aphid Myzus persicae. (a) anatomical features; (b) organization in intact animal. (Redrawn from Fig. 5 of Ponsen (1979) and Fig. 2.1.6F of Ponsen (1987).)
into the midgut (this valve prevents the back flow of food from the midgut when the pharyngeal pump dilates (see Section 3.1)). The midgut comprises a proximal ‘stomach’, which is dilated or tubular, and a distal ‘intestine’, and it lacks any caeca or other diverticula. The transition from midgut to proximal hindgut is not marked by Malpighian tubules, which are absent from aphids, or any other anatomical feature. The distal portion of the hindgut is the rectum, a thin-walled, distensible sac in which egesta accumulate prior to expulsion as a single drop of honeydew, every 15–45 min. The aphid gut is 2–4 times longer than the body, varying between aphid species. The intestine (i.e. distal midgut) contributes much of the length of the aphid gut and it is looped several times within the body cavity. The oesophagus and hindgut include no loops or coils. In many aphids, including most members of the Aphidinae, the stomach is closely apposed to more distal regions of the gut. In some aphids, notably members of the subfamilies Callaphidinae, Lachninae and Drepanosiphinae, this anatomical association is elaborated, such that a portion of the midgut is encircled by a more distal region of the alimentary tract, an arrangement described as a filter system.
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(Contrary to early suggestions in the literature, the proximal region is not completely enclosed within the distal region, and the two regions are separated by a narrow haemolymph-filled space (Ponsen, 1981).) For example, in Eulachnus brevipilosis and other lachnines, the most proximal region of the midgut is enclosed within the proximal hindgut; and the callaphidine Subsaltusaphis ornata has two filter systems, a first in which the stomach is encircled by the proximal hindgut and a second involving very close apposition between the proximal and distal intestine (Ponsen, 1979, 1981). The close apposition between proximal and distal regions of the alimentary tract and particularly the filter systems in certain species is reasonably interpreted as facilitating the transfer of compounds between different regions of the gut, as is discussed in Section 4.3.3. The gut wall in aphids comprises a single layer of epithelial cells bounded externally by the basal lamina and musculature. The latter is limited to scattered circular muscle cells for the foregut and midgut, and a well-developed circular muscle plus scattered longitudinal muscle fibres for the hindgut (Ponsen, 1987). The poorly developed gut musculature is consistent with the food being forced through the gut principally by proximal pressure exerted by the plant sieve elements or aphid cibarial pump, with the possible assistance of peristaltic movements of the hindgut. The gut epithelial cells are separated from the gut lumen by a chitinous cuticle for the foregut and hindgut, as in other insects. At each moult of the insect, the cuticle is shed; the oesophageal cuticle is passed back into the stomach, where it remains through the life of the insect. The midgut of aphids and other hemipteran insects lacks a peritrophic matrix and the apical surface of the midgut cells is bounded by a continuous sheet of lipoprotein that Terra (1990) call the perimicrovillar membrane. The perimicrovillar membrane of aphids has not been studied in detail, but is probably similar in origin and general organization to that in other hemipteran insects, e.g. Silva et al. (1995). The structure of the epithelial cells of the aphid midgut has been investigated in detail, especially by M.B. Ponsen (reviewed in Ponsen, 1987). Consistent with the role of the midgut as the site of enzyme production and nutrient assimilation, the apical cell membrane of midgut cells is elaborated into microvilli or, especially in the stomach, more complex structures that Ponsen terms ‘microlabyrinths’. The basolateral membranes of many cells are also highly folded, indicative of a role in transport. The early literature (summarized in Ponsen, 1987) concluded that digestive enzymes are secreted into the aphid gut lumen by apocrine secretion, i.e. the shedding of portions of cytoplasm by cellular constriction or budding. Profiles of cells with the predicted apical extensions are evident in the stomach of sectioned aphids prepared by traditional histological methods. However, the contribution of fixation artefacts to these observations cannot be excluded definitively (Brunings and DePriester, 1971) and the significance of apocrine secretion to aphid gut function deserves re-examination with modern microscopical techniques.
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Recent studies implicate the perimicrovillar membrane as an important site of enzyme activity, as is considered in detail for sucrase in Section 4.3.1.
4.3
PROCESSING OF INGESTED SUGARS
Virtually all research to date on the fate of sugars ingested by aphids has concerned dietary sucrose. Although oligosaccharides of the raffinose series and polyols are important, phloem compounds in many plants (see Section 2.2.1), their utilization by aphids has not been investigated, beyond the early demonstration of F. Dupsiva (quoted in Srivastava, 1987) that homogenates of certain aphid species can hydrolyse raffinose to its constituent monosaccharides.
4.3.1
Evidence from honeydew sugars
Much of the sucrose ingested by aphids is not assimilated and is voided via the honeydew (e.g. see Fig. 6). However, the ingested sucrose is undetectable or accounts for a very small proportion of the honeydew sugars, suggesting that the voided sugars are the product of extensive enzymatic modification in the insect gut. Auclair (1963) reviewed the extensive early literature showing that the honeydew of various aphid species may contain the trisaccharide melezitose [Glc-(1-3)-Frc-(2-1)-Glc], the tetrasaccharide maltosucrose [Glc-(1-4)-Glc(1-4)-Glc-(1-2)-Frc] and higher order oligosaccharides; and these conclusions have been confirmed amply by subsequent studies (e.g. Walters and Mullin, 1988; Hendrix et al., 1992; Wilkinson et al., 1997). Aphid honeydew may also contain isomers of sucrose, especially trehalulose [Glc-(1-1)-Frc] and turanose [Glc-(1-3)-Frc]. For aphids reared on chemically defined diets of different sucrose concentrations, the contribution of oligosaccharides to honeydew sugars increases with dietary sucrose concentration (Fig. 8a), and the dominant or sole sugar in these oligosaccharides was glucose (Ashford et al., 2000). Additional information comes from experiments in which pea aphids were fed on diets with sucrose radioactively labelled in the glucose or fructose moiety and the distribution of radioactivity in honeydew components was quantified. Consistent with the high glucose content of honeydew oligosaccharides (see above), the radioactivity recovered from the honeydew of aphids fed with [14C-glucose]-sucrose was approximately a hundred-fold higher than for those fed with [14C-fructose]-sucrose (Ashford et al., 2000). These data indicate that the glucose moiety of much of the ingested sucrose is channelled into oligosaccharide synthesis and voided via the honeydew, while the sucrose-derived fructose is assimilated quantitatively from the gut into the aphid tissues.
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FIG. 8 Impact of sucrose concentration on sugar transformations in the pea aphid Acyrthosiphon pisum. (a) The contribution of oligosaccharides to honeydew produced by aphids reared on chemically defined diets; (b) the production of glucose by gut homogenates. The observed rate differs from the predicted rate curve for a reaction following saturation kinetics, which is shown as a broken line. (Redrawn from Fig. 2 of Wilkinson et al. (1997) and Fig. 2a of Ashford et al. (2000).)
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Evidence from gut enzymes
Crude homogenates of guts dissected from aphids hydrolyse sucrose to its constituent monosaccharides at a high rate. The sucrase is a -glucosidase, i.e. it releases -D-glucose from the non-reducing end of -linked substrates (Srivastava and Auclair, 1962; Ashford et al., 2000). The crude gut homogenates also display appreciable activity against trehalose and melezitose, but very low activity against isomaltose. The number of enzymes contributing to these capabilities is unknown, but the aphid enzyme differs from the mammalian intestinal sucrase, which is a bifunctional isomaltase–sucrase. Fragmentary and partly inconsistent information is available on the localization of the aphid sucrase. In pea aphids, sucrase activity has been detected in saliva (Miles, 1999) and the midgut, but with contradictory reports of the restriction of the midgut enzyme to the stomach (Rhodes et al., 1997) and intestine (Ashford et al., 2000). The doubling in sucrose activity by addition of detergent to homogenates (Ashford et al., 2000; Cristofoletti et al., 2003) has been interpreted as evidence that the enzyme is bound to the perimicrovillar membrane (see Section 4.2), possibly on the luminal face, as for the -glucosidase of the heteropteran Dysdercus peruvianus (Silva et al., 1995). The reasoning is that, on homogenization, the perimicrovillar membrane, but not the gut epithelial cell membrane, would generate inside-out and rightside-out vesicles in equal proportions and enzyme activity on the inside of the vesicles would not have access to substrate in the absence of detergent treatment. Anchored to the perimicrovillar membrane, the sucrase would not be lost from the midgut with the stream of food. Consistent with this interpretation, sucrase activity is undetectable in aphid honeydew. Crude gut homogenates of several aphid species also mediate the synthesis of oligosaccharides from sucrose (Duspiva, 1955). In homogenates of pea aphid guts incubated with sucrose as substrate, the production of glucose and fructose increases with sucrose concentration up to 50 mM sucrose, but glucose production declines at higher sucrose concentrations (Fig. 8b) (Ashford et al., 2000). It has been suggested that a single enzyme may mediate both sucrose hydrolysis and oligosaccharide synthesis, by inserting a molecule of water and glucose, respectively, at the glucosidic bond (Walters and Mullin, 1988; Ashford et al., 2000). The synthesis of oligosaccharides by this reaction is called transglucosidation. Indeed, it is possible that the wide array of sugars reported in aphid honeydew (see Section 4.3.1) is generated by a single enzyme. For comparison, the bacterium Neisseria polysaccharea possesses an enzyme informally known as an amylosucrase that transforms sucrose into a diversity of products, including glucose, fructose, sucrose isomers, malto-oligosaccharides and long-chain oligosaccharides (Montalk et al., 2000). A candidate sucrase is an -glucosidase of
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molecular weight 70 kDa with sucrase activity, purified from the pea aphid (Cristofoletti et al., 2003). However, the sucrase–transglucosidase activity of aphids may be mediated by multiple isozymes with slightly different kinetic properties.
4.3.3
How chemical transformations of sucrose in the gut contribute to aphid osmoregulation
The osmotic pressure of phloem sap is 2–4 times greater than that of aphid body fluids. As a result, an aphid would be expected to lose water to the lumen of the alimentary tract and literally shrivel as it fed. Osmoregulatory collapse is circumvented by downregulation of the osmotic pressure of the gut contents, such that the aphid honeydew and haemolymph are isosmotic. As an example, pea aphids (A. pisum) feeding from their host plant Vicia faba imbibe phloem sap of osmotic pressure 2 MPa and produce honeydew of osmotic pressure 0.9 MPa, equal to that of their haemolymph (Wilkinson et al., 1997). Sugar transformations in the alimentary tract have been implicated in aphid osmoregulation. The transformation of ingested disaccharides to oligosaccharides would tend to reduce the osmotic pressure of the gut contents because the osmotic pressure exerted by solutes is determined by the molality of the solutes, and not their weight. However, the sucrose concentration at which oligosaccharides are synthesized in vivo (>0.3 M, see Fig. 8a) is markedly higher than that predicted from experiments in vitro (>0.05 M) (Fig. 8b) (Wilkinson et al., 1997; Ashford et al., 2000), raising the possibility that the ingesta are diluted prior to enzymatic transformation. The anatomical basis for such a process may be the looping of the distal intestine in close apposition to the stomach (see Section 4.2). If, as argued by Ashford et al. (2000), the sucrase/transglucosidase is localized to the perimicrovillar membrane of the intestine, then the osmotic pressure in the distal intestine would be reduced relative to the stomach, and water would pass down its osmotic gradient from intestine to stomach (Fig. 9), as also proposed by Rhodes et al. (1997). Water channels at the sites of intestine–stomach apposition may promote water flux. By this scenario, the uncontrolled movement of water from haemolymph to stomach would be reduced by two factors: first, the lowered osmotic differential between the two compartments, arising from the dilution of stomach contents; and, second, the hydrostatic pressure of the ingesta, generated in the plant sieve tubes or by the aphid cibarial pump and maintained in the relatively inextensible stomach chamber. Although transglucosidation of ingested sugars in the gut can substantially reduce the osmotic pressure of the gut contents, it cannot be the exclusive basis of aphid osmoregulation, such that the honeydew osmotic pressure equals that of the haemolymph (see above). This is because the transglucosidase activity
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FIG. 9 Contribution of water flux to osmoregulation of aphids. It is proposed that the osmotic pressure in the distal intestine is lower than that in the stomach, as a result of oligosaccharide synthesis in the proximal intestine, resulting in the net passage of water (shown as the triple arrow) from distal intestine to stomach. (Redrawn from Fig. 2.9c of Dixon (1998).)
varies with sucrose concentration and not with the total osmolality of the solution. The transglucosidase activity in the gut lumen is probably complemented by the transport of water from the gut to the haemolymph, linked to active ion transport. The site(s) of water transport is unknown. In principle, the rectum that, in other insects, plays a vital role in water reabsorption is a likely candidate, especially as the ejecta accumulate in the rectum prior to elimination as honeydew. The rectal epithelium of aphids is very thin, offering minimal barrier to water and ion movement and the basal cell membrane has elaborate folding, characteristic of ion and water transporting tissues. Current understanding of the relationship between sugar transformations and water flux in the aphid alimentary tract is the reverse of the perception in the early literature. For example, Goodchild (1966) and House (1974) argued that water and sugars are transferred from the stomach to the hindgut, bypassing the intestine where nutrients are assimilated. As considered above, the likely direction of passage of water is from the distal midgut/ proximal hindgut to the stomach in aphids and other phloem-feeding insects (see Fig. 9).
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FIG. 10 Contribution of essential amino acids (open symbols) and non-essential amino acids (closed symbols) to the honeydew of aphids reared on a diet containing essential and non-essential amino acids in 1:1 ratio. (Redrawn from data in Fig. 2b of Prosser et al. (1992).)
4.4 4.4.1
PROCESSING OF INGESTED NITROGENOUS COMPOUNDS
Amino acids
The principal nitrogenous compounds in the phloem sap of most plants are free amino acids (Section 2.3), and these compounds can be assimilated without prior modification across the gut wall of animals. It is evident from the early literature that the overall amino acid assimilation efficiency is high, e.g. 55% for the willow aphid Tuberolachnus salignus feeding from Salix twigs (Mittler, 1958), and 64–70% for M. persicae on chemically defined diets (Kunkel and Hertel, 1975). One might anticipate that aphids would preferentially take up essential amino acids, the 9 of the 20 amino acids that contribute to protein and cannot be synthesized by animals. Very commonly, however the assimilation efficiency for non-essential amino acids is higher than for essential amino acids for aphids feeding from both chemically defined diets and plants; the illustrative data in Fig. 10 for A. pisum on diets is indicative of a consistent differential of ca. 10 mM between the assimilation of essential and nonessential amino acids. The reason for this apparently anomalous result is that the aphid requirement for essential amino acids is met, at least in part, by their symbiotic bacteria (see Section 5). Because the essential amino acid content of phloem sap is low, the preferred assimilation of non-essential amino acids by aphids can result in honeydew with a more balanced amino acid composition than in phloem sap. For example, the essential:non-essential amino acid ratio for the phloem sap of V. faba is 1:10, and of the honeydew of aphids feeding on this plant is 1:2 (unpublished data). The transport of amino acids across the gut epithelial cells to the haemolymph of aphids has not been investigated directly but there is a strong
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expectation that, as in other insects, it is mediated by cation-coupled transporters, possibly complemented by high-capacity uniporters, on the apical membrane of midgut cells (Wolfersberger, 2000). Multiple transporters are anticipated, at least some with broad but overlapping amino acid specificities. Most research on amino acid transport in other insects has concerned the transport of neutral amino acids, especially leucine, and there is excellent evidence that leucine uptake is coupled to inward-directed K þ flux at the apical membrane of midgut cells in phytophagous Lepidoptera and Coleoptera (e.g. Castagna et al., 1998). The recruitment of K þ as the symport ion (in contrast to Na þ used by most animals) is seen as an adaptation to the high K þ content and K þ :Na þ ratio of plant foodstuffs and, consequently, of the gut lumen content of phytophagous insects. If this reasoning is correct, one might also expect the aphid gut transporters to be K þ -linked, given the high K þ concentration in phloem sap (see Section 2.2.4). The mechanism of amino acid efflux from midgut cells to the haemolymph has not been characterized in insect systems. Terra (1990) has constructed a detailed model to account for the efficient uptake of amino acids from the gut lumen into the midgut epithelial cells of Hemiptera, including aphids. The core element to this model is a proposed low concentration of K þ ions in the space between the perimicrovillar membrane and midgut epithelial cells, generated by active transport of K þ ions across the apical membrane of the gut epithelial cell (Fig. 11). The resultant concentration gradient of K þ ions across the perimicrovillar membrane provides the driving force for K þ -coupled transport of amino acids across this membrane. The amino acids that accumulate in the perimicrovillar space are then envisaged to diffuse to carriers in the microvillar membrane, by which they are translocated into the cells. To the author’s knowledge, the
FIG. 11 Model for amino acid uptake from the gut lumen into a gut epithelial cell. Transporters are indicated as circles: the amino acid-K þ symport on the perimicrovillar membrane, and the ATPase-driven active transport of K þ ions and facilitated diffusion of amino acids into the epithelial cell. (Redrawn from Fig. 8.4b of Terra et al., 1996.)
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validity of this scheme has not been investigated experimentally for aphids or any other hemipteran insect. 4.4.2
Proteins
The presence of proteins in plant phloem sap, albeit usually at low concentrations, raises the possibility that aphids may degrade ingested proteins to amino acids, as a supplementary source of nitrogen. To date, virtually all the research has been done on A. pisum, the host plants of which are legumes with low phloem protein content. Aphids feeding from cucurbits, which have protein-rich phloem sap, have not been studied. Analyses of gut homogenates of A. pisum have revealed aminopeptidase activity, especially in the distal midgut, but no endoprotease activity (Srivastava and Auclair, 1963; Rahbe´ et al., 1995). Consistent with these data, Rahbe´ et al. (1995) demonstrated that aphids feeding from chemically defined diets supplemented with various lectins produce honeydew containing these proteins, apparently without any modification. There are, however, two limitations to these studies. First, the gut epithelium is not an absolute barrier to proteins, which may be translocated to the haemolymph (Down et al., 2000). The mode of transfer of these proteins is obscure (they may pass between epithelial cells) but, once in the haemolymph, they would be available for degradation by haemolymph proteases. The incidence of haemolymphmediated degradation of ingested proteins and its significance to aphid nutrition have not been investigated. Second, the fate of proteins naturally occurring in plant phloem sap and ingested by aphids may differ from that of lectins studied by Rahbe´ et al. (1995); lectins are generally rather resistant to proteolytic cleavage. The value of examining the proteolytic capability of aphid alimentary tracts further is indicated by some fragmentary data that other phloemfeeding homopteran insects may degrade ingested protein. Guts dissected from the rice brown planthopper Nilaparvata lugens contain appreciable trypsin-like and cathepsin B-like protease activity (Foissac et al., 2002), although the activity of these enzymes against ingested proteins remains to be tested. The whitefly Bemisia argentifolii fed on radioactively labelled leaf proteins generates honeydew containing low molecular weight radioactive compounds (Salvucci et al., 1998), but it is not known whether the honeydew compounds were amino acids, whether the ingested proteins were hydrolysed in the insect gut and, if so, why the resultant amino acids were not assimilated into the tissues. 4.5
THE FATE OF INGESTED ALLELOCHEMICALS
Phloem-mobile secondary compounds are effective defences against many aphid species. As examples, when M. persicae feeds from the castor oil plant
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Ricinus communis, the phloem-mobile alkaloid ricinine accumulates rapidly in the aphid tissues and the aphids die within 8–24 h (Olaifa et al., 1991); and lines of the lupin Lupinus angustifolius with high alkaloid levels in their phloem sap are more resistant to M. persicae than lines with low phloem alkaloid levels (Berlandier, 1996). However, chemical defences are not universally effective against phloem-feeding insects. Some aphid species detoxify the compounds, eliminate them via the honeydew or sequester them in their tissues (Dixon, 1998) and the sequestered compounds can additionally confer protection from predators (e.g. Rothschild et al., 1970). However, the interaction between these compounds and processes in both the gut and wider aphid tissues are poorly understood at the physiological and biochemical levels. Current understanding of three systems that have been the subject of sustained research are considered here. 4.5.1
Pyrrolizidine alkaloids
The principal plants used to study the impact of pyrrolizidine alkaloids on insects are composites of the genus Senecio, especially S. jacobea which has a single phloem-mobile pyrrolizidone alkaloid, senecionine-N-oxide (Hartmann and Dierich, 1998). Senecionine-N-oxide is not toxic but it is transformed into an array of highly reactive and toxic alkylating agents by the activity of insect cytochrome P450 enzymes (Hartmann, 1999). These reactions are an ‘Achilles heel’ of insect cytochrome 450s, which function generally to degrade plant allelochemicals and other xenobiotics to non-toxic products (Schuler, 1996). The aphid Aphis jacobaea is a specialist on Senecio species and it avoids the toxicity of the alkaloid by its possession of a flavin monoxygenase, which efficiently transforms senecionine back to the non-toxic N-oxide (Hartmann, 1999). The concentration of pyrrolizidine alkaloids in the aphid tissues is ca. 20 mg g 1 dry weight, an order of magnitude lower than both its honeydew and feeding plant, suggesting that senecionine-N-oxide is also selectively eliminated via the honeydew. Even so, A. jacobaea preferentially occurs on S. jacobea plants of low alkaloid content, suggestive of limits or costs to the avoidance of alkaloid poisoning in this aphid (Vrieling et al., 1991). 4.5.2
Glucosinolates
Crucifer-feeding aphids process glucosinolates, the dominant defensive compounds in these plants, without ill effect. Glucosinolates are one component in a plant ‘binary chemical weapon’; the other component is an enzyme, generically known as myrosinase, localized in different plant cells from the glucosinolates (Kelly et al., 1998; Korslova et al., 2000), such that the myrosinase and glucosinolates come into contact only when the plant is damaged, for example by a chewing herbivore or necrotrophic pathogen. At that point, the myrosinase hydrolyses the glucosinolate to an unstable
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aglycone, which undergoes spontaneous re-arrangement to yield various toxic compounds, including isothiocyanates, thiocyanates and nitriles. The phloem-mobile glucosinolates (see Section 2.2.5) are not hazardous to aphids because aphid feeding does not disrupt the compartmentation of the plant glucosinolates and myrosinase, and ingested glucosinolates may either pass directly through the aphid alimentary tract to the honeydew or accumulate, apparently unaltered, in the aphid tissues, probably the haemolymph. There is evidence that the fate of gluosinolates varies between aphid species, with preferential accumulation in the specialist crucifer-feeding aphids, Brevicoryne brassicae and Lipaphis erysimi, and preferential elimination via the honeydew in M. persicae, a polyphagous aphid which utilizes both crucifers and noncruciferous plants (e.g. Merritt, 1996; Bridges et al., 2002). The specialist species also differ from M. persicae in that the former contain readily detectable myrosinase activity (MacGibbon and Allison, 1968), localized to microbodies in the skeletal muscle (Bridges et al., 2002). The myrosinase enzyme of B. brassicae has been purified. It is a homodimer of 54 kDa (Jones et al., 2001) and its gene sequence displays greater similarity to animal -Oglucosidases than to plant myrosinases (Jones et al., 2002). Since the toxicity of glucosinolates is mediated by myrosinase activity, aphid possession of myrosinase is, at first sight, counter-intuitive. However, Jones et al. (2001) have argued that glucosinolate hydrolysis to toxic products in an aphid damaged by a natural enemy would deter the enemy from attacking other members (presumably clone-mates) of an aphid colony, even though the victim would be unlikely to survive an attack sufficiently damaging to disrupt the glucosinolate–myrosinase compartmentation. The implication is that the myrosinase–glucosinolate binary defence has evolved independently in plants and aphids. However, the significance of sequestered glucosinolates as an aphid defence remains to be quantified. 4.5.3
The condition of the stomach in aphids feeding from chenopods and other plants
When aphids feed on plants of the family Chenopodiaceae (e.g. sugarbeet, fat hen), their stomachs swell to linear dimensions up to double that of aphids of similar size on non-chenopods, and become filled with a white precipitate (Edwards, 1966), which subsequently gives rise to a black deposit. The impact on aphid performance is variable; the early decline of M. persicae on sugarbeet crops has been linked to the stomach precipitate (Williams, 1995), but Aphis fabae performs well on Chenopodium album, despite their inflated stomach (Wilkinson et al., 2001). The white precipitate comprises a polymer of glucose and amino acids (Williams et al., 1997), raising the possibility that gut processing of ingested sugars and amino acid metabolism may be affected deleteriously. However, the nature of the stomach malfunction and the component of chenopod phloem sap that induces it are unknown.
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Chenopods are not unique in their impact on the organization on the stomach of aphids. The stomach of aphids feeding from crucifers has been reported to accumulate rod-shaped inclusions of length up to 0.3 mm. The chemical composition of these structures is unknown, but the experiments of Moericke and Mittler (1966) indicate that they are formed in the oesophagus and passed back to the stomach, where they are retained for the remainder of the insect’s life. These inclusions appear to have no impact on the aphid performance, and Moericke and Mittler (1966) suggest that they may be a component of the aphid saliva that have been modified or precipitated by unknown compounds in the phloem sap of crucifers. 5
Acquisition of nutrients from symbiotic micro-organisms
A central feature of the nutritional physiology of aphids is that these insects obtain their nutritional requirements from two sources: their food and their symbiotic micro-organisms. The microbiota of most members of the Aphididae (see Fig. 1) is dominated by a single clade of bacteria, -proteobacteria assigned to the genus Buchnera, known only in aphids (Munson et al., 1991). Aphids of the tribe Cerataphini have yeasts, and not Buchnera. The microbiota in the Adelgidae and Phylloxeridae has not been studied in recent years, but it apparently does not include Buchnera (Buchner, 1965). Over the last decade, research on the aphid–Buchnera symbiosis has been dominated by the molecular biological and genomic studies, including the complete genome sequences of Buchnera from A. pisum (Shigenobu et al., 2000) and S. graminum (Tamas et al., 2002). This section provides a brief overview of the microbiology of aphids (Section 5.1), followed by more detailed consideration of the role of the symbiosis in aphid nutrition. 5.1
THE MICROBIOLOGY OF APHIDS
In most insects, the greatest abundance and diversity of micro-organisms occur in the lumen of the alimentary tract, and relatively few microbial taxa breach the gut wall to the internal organs. Aphids are unusual in that the aphid alimentary tract bears few or apparently no micro-organisms detectable by microscopy or molecular methods (Grenier et al., 1994; Harada et al., 1996; Wilkinson et al., 1997). This is probably a consequence of the highly disturbed condition in the aphid gut, which approximates to a simple tube (lacking Malpighian tubules, midgut caeca or any other diverticula, see Section 4.2) through which the liquid diet flows in a single direction. Micro-organisms ingested with the food generally fail to persist in the alimentary tract of aphids (Harada and Ishikawa, 1997). Cells of Buchnera sp. dominate the microbiology of aphids. They occur at a density of ca. 107 cells mg 1 aphid fresh weight, equivalent to ca. 10% of the total volume of the insect (Baumann et al., 1994; Humphreys and Douglas,
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1997; Wilkinson et al., 2001). The aphid–Buchnera symbiosis has three defining features: (1) The Buchnera cells are intracellular. They are located in a single cell type, known as the mycetocyte or bacteriocyte, the sole function of which is apparently to maintain the bacteria which occupy 60–70% of the mycetocyte cytoplasm (Whitehead and Douglas, 1993a). Each Buchnera cell is surrounded by a membrane of insect origin, known as the symbiosomal membrane. The mycetocytes are located in the haemocoel of the insect, forming a coherent V-shaped organ dorsal to the alimentary tract, with the base of the ‘V’ towards the posterior end of the aphid body. Mycetocytes differentiate in the early aphid embryo. Consequently, parthenogenetic aphids, which bear embryos in their ovaries, have mycetocytes in both the maternal haemocoel and in the haemocoel of all but the youngest embryos. In an adult aphid, up to 75% of the Buchnera population is in the embryos (Humphreys and Douglas, 1997). (2) The Buchnera cells are invariably transmitted vertically from mother aphid to her offspring via the ovaries. The bacterial cells are exocytosed from maternal mycetocytes abutting the germarium and pass between the ovarial follicle cells to be incorporated into the unfertilized egg of oviparae (aphid females which produce sexual eggs) or blastoderm embryo in virginoparous aphids (Hinde, 1971; Brough and Dixon, 1990). The fidelity of vertical transmission is indicated by the remarkable congruence between the phylogenetic trees of Buchnera and of their aphid hosts, constructed from gene sequence data (Moran et al., 1993). These data have, further, been used to estimate the date of origin of the symbiosis, at 180–250 million years ago, a timescale compatible with the likely time of evolutionary origin of the aphids. (3) The association is obligate for both the aphid and Buchnera. Buchnera can be isolated from aphids and persists for several hours (Whitehead and Douglas, 1993b), but it cannot be maintained indefinitely apart from the insect. Buchnera has a very small genome size, varying between 0.45 and 0.64 Mb (Shigenobu et al., 2000; Wernegreen et al., 2000; Gil et al., 2002; Tamas et al., 2002), and lacks many genes required for an independent existence, including various genes in intermediary metabolism, nutrient translocation, signal exchange and DNA repair (Shigenobu et al., 2000; Silva et al., 2001; Tamas et al., 2002). The requirement of aphids for their bacteria is indicated by their poor growth and collapse in their reproductive output when the Buchnera are eliminated with antibiotics (Mittler, 1971; Douglas, 1992). The bacteria-free aphids generated with antibiotics are known as aposymbiotic aphids. Overall, Buchnera cells account for 90% or more of the total number of microbial cells in an aphid. Among the other micro-organisms are bacteria known as ‘secondary symbionts’ or ‘accessory bacteria’ that are vertically transmitted along with the Buchnera. They differ from Buchnera in that they have a wider tissue distribution, often occurring in haemolymph and sometimes associated with the cells lining the alimentary tract, they can be transmitted horizontally, and they are not universal in natural populations of
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some aphid species (Fukatsu et al., 2000; Darby et al., 2001; Sandstro¨m et al., 2001). These bacteria have been linked to various attributes of the aphids, including enhanced tolerance of elevated temperature (Montllor et al., 2002) and plant range (Chen et al., 2000), but the physiological basis of these interactions has not been investigated.
5.2 5.2.1
THE NUTRITIONAL CONTRIBUTION OF BUCHNERA TO APHIDS
Production of essential amino acids by Buchnera
There is overwhelming evidence that aphids derive essential amino acids from their complement of Buchnera cells. The essential amino acids are the 9 amino acids which contribute to protein but cannot be synthesized de novo by animals, as distinct from the 11 amino acids in protein that animals can synthesize. An endogenous supply of essential amino acids is crucial to aphids because the ratio of essential amino acids:non-essential amino acids is generally 1:3 to 1:10 in plant phloem sap (see Section 2.2.3), and 1:1 in aphid protein, which accounts for >90% of the total amino acid content of the insect. The first direct evidence for bacterial provisioning of essential amino acids came from the study of Mittler (1971) on the green peach aphid, M. persicae, maintained on chemically defined diets from which each of the 20 amino acids were individually omitted. All of the diets except the methionine-free diet supported sustained aphid growth, with the implication that M. persicae is independent of a dietary supply of most amino acids. Mittler (1971) attributed the requirement for methionine to the phagostimulatory properties of this amino acid, and not to an absolute dietary requirement. A parallel group of aphids was treated with the antibiotic aureomycin ( ¼ tetracycline) to eliminate the bacteria, and these insects required all essential amino acids, as expected if the bacteria provided the essential amino acids. The growth rate of aphids on diets with individual essential amino acid omissions has been used subsequently to quantify the net rates of amino acid production by Buchnera (Douglas et al., 2001). The amino acid composition of aphid protein is relatively invariant and therefore the rate at which each amino acid is incorporated into protein can be calculated from the overall protein growth rate of the insect. If an amino acid is omitted from the diet, it is incorporated into aphid protein at a rate determined by the rate of provisioning by the Buchnera cells and its availability in non-protein reserves (e.g. the free amino acid pool in the haemolymph) for aphids containing Buchnera, and at a rate determined by the pre-existing reserves for aposymbiotic aphids. Consequently, the net rate of Buchnera-derived provisioning can be quantified from the difference in protein growth rates between aphids containing and lacking their bacteria. Representative data for each of the essential amino acids in the black bean aphid A. fabae (Table 1)
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TABLE 1 Net synthesis of essential amino acids by Buchnera in Aphis fabae reared on diets lacking individual essential amino acids (Data from Table 2 of Douglas et al. (2001).) Mean rate of protein-amino acid synthesis (pmol g 1 total protein day 1) Amino acid Histidine Isoleucine Leucine Lysine Methionine Phenylalanineb Threonine Tryptophan Valine
Rate of protein-amino acid synthesized by Buchnera
symbiotic aphidsa
aposymbiotic aphidsa
pmol g 1 total protein day 1
fmol Buchnera cell 1 day 1
12.4 114.7 180.8 157.4 31.6 84.4 111.3 10.3 128.3
3.3 21.9 24.7 32.1 13.1 – 19.3 2.6 22.6
9.1 92.8 156.1 125.3 18.5 – 92.0 7.7 105.7
0.02 0.19 0.33 0.26 0.04 – 0.19 0.02 0.22
a
Symbiotic aphids bear their normal complement of symbiotic bacteria, and aposymbiotic aphids were treated with the antibiotic rifampicin to eliminate the symbiotic bacteria. b The protein growth rate of aposymbiotic aphids on phenylalanine-free diet could not be quantified because of high mortality.
indicate that the amino acids are produced at net rates varying between 0.02 and 0.33 fmol bacterial cell 1 day 1. Consistent with these nutritional studies, aphids containing their normal complement of Buchnera can synthesize essential amino acids. For example, when A. pisum is fed with 14C-sucrose, radioactivity is recovered from all of the protein amino acids, including the essentials, but incorporation into the essential amino acids is abolished in aposymbiotic aphids (Febvay et al., 1999). Other studies have demonstrated the metabolism of 14C-glutamate to the essential amino acids isoleucine, lysine, methionine and threonine in A. pisum (Febvay et al., 1995) and A. fabae (Douglas et al., 2001; Wilkinson et al., 2001), 14 C-anthranilate to tryptophan in A. pisum (Birkle et al., 2002), and 35SO4 to methionine in M. persicae (Douglas, 1988). These results have received excellent confirmation from the complete genome sequence data of Buchnera from A. pisum. Although the genome is small and lacking in many metabolic capabilities, including the capacity to synthesize all non-essential amino acids apart from cysteine, the genes coding for virtually all enzymes in the synthesis of the nine essential amino acids are present. The most plausible explanation for this condition is that the evolutionary processes promoting gene loss in Buchnera (reviewed in Rispe and Moran, 2000) are countered by selection to retain the capacity to synthesize essential amino acids. There are, however, two potentially important exceptions to this generality. Annotation of the genome of Buchnera from A. pisum and S.
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graminum indicates that ilvA, coding for threonine deaminase the first enzyme in the dedicated isoleucine biosynthetic pathway, and metC, coding for cystathione -lyase in the methionine biosynthetic pathway, are absent; but nutritional and metabolic data suggest that various aphids, including A. pisum, can synthesize both isoleucine and methionine. Possible explanations for the discordance between the genomic and other data have been considered in detail in Douglas et al. (2002). Briefly, enzymes other than threonine deaminase and cystathione -lyase may catalyse the reactions in question through low substrate specificity; the accessory bacteria (see Section 5.1) may mediate these reactions; or the genes ilvA and metC may have been transferred to the aphid genome and transcribed and translated in the nucleocytoplasm of the mycetocytes with transfer of the protein to the Buchnera cells. 5.2.2
Integration of Buchnera production of essential amino acids into aphid nutritional physiology
Aphid demand for Buchnera-derived essential amino acids is anticipated to vary, for example with phloem sap amino acid composition and content and with aphid developmental age or physiological condition. One might expect that the rate of amino acid production and profile of amino acids produced by Buchnera cells would vary according to aphid demand, such that Buchnera nutritional function is integrated into the wider nutritional physiology of the aphid. However, the genomic data of Buchnera can be interpreted to offer no support for this expectation. Amino acid synthesis in bacteria is regulated by a complex network of controls over transcription rates and enzyme activity, principally mediated by feedback repression and inhibition, respectively, by the amino acid end product. In this way, an amino acid is synthesized at high rates only when it is in short supply. In Buchnera, however, transcriptional regulators, including attenuation systems, are apparently absent for all genes contributing to amino acid synthesis (Shigenobu et al., 2000; Tamas et al., 2002). The implication is that essential amino acid production by Buchnera is unlikely to be product-regulated. An alternative scenario is that essential amino acid production by Buchnera may be controlled by the supply of precursors of the amino acid biosynthetic pathways. Because Buchnera cells are intracellular, they, of necessity, derive all their nutrients, including precursors of essential amino acid synthesis, from the surrounding insect cell cytoplasm. However, Buchnera cells have very limited capacity for the selective uptake of compounds. The genomic data indicate that Buchnera has very few transporter genes, specifically a few ABC transporters, low-affinity transporters and porins which promote passive diffusion (Shigenobu et al., 2000). The supply of nutrients to Buchnera is therefore probably determined by the transport capacity of the insect symbiosomal membrane which surrounds each Buchnera cell. In principle, the aphid could control the synthesis of essential amino acids in Buchnera by regulated
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supply of precursors across the symbiosomal membrane to each Buchnera cell. At present, there are no experimental data to assess the validity of such a scenario and the properties of aphid symbiosomal membrane are completely unknown. Metabolic approaches have, to date, contributed little to our understanding of the response of the symbiosis to variation in aphid demand for essential amino acids. The principal approach used has been to explore the impact of dietary supply of amino acids on the incorporation of radioactivity into essential amino acids (Febvay et al., 1999; Douglas et al., 2001). When aphids are reared on diets from which an essential amino acid is omitted or provided at reduced concentration, they incorporate radioactivity from a radiolabelled precursor into that amino acid at elevated rates relative to aphids feeding from a diet with higher concentration of the amino acid. It has been concluded from these experiments that the absolute rate of amino acid synthesis and profile of amino acids produced by Buchnera cells are not fixed, but may vary according to aphid demand. However, this interpretation may generally be faulty because it does not take into account the specific activity of the radiolabelled precursor at the site of essential amino acid synthesis. In other words, the radioactivity in an amino acid synthesized by aphids on a diet lacking that amino acid may be elevated because the absolute concentration of the amino acid is depressed rather than because its rate of synthesis is elevated. To resolve this issue, future experiments are needed to address the total flux of carbon/nitrogen through the amino acid biosynthetic pathways in Buchnera.
6
Fate of nutrients acquired by aphids
The fate of nutrients acquired by aphids from their diet and bacterial symbiosis can be considered from two perspectives: the pattern of allocation of nutrients to the various chemical classes of compounds, as shaped by flux through different metabolic pathways; and the allocation of nutrients to different organ systems, resulting in the partitioning of resources between maintenance, growth, reproduction etc. These two perspectives, the metabolic and physiological fate of nutrients, respectively, are considered in turn. 6.1 6.1.1
METABOLIC FATE OF ACQUIRED NUTRIENTS
Carbohydrates
Early studies revealed that the respiratory quotient of various aphid species approximates to unity, indicating that carbohydrate is the dominant respiratory substrate (summarized in Table 2 of Rhodes et al., 1996). This conclusion has been confirmed amply by recent respirometry analyses (Salvucci
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TABLE 2 Metabolic fate of sucrose ingested by Acyrthosiphon pisum from a chemically defined diet containing 0.58 M sucrose supplemented with 14C-sucrose and 0.26 M amino acids of composition mimicking phloem sap. (Approximately 23% of assimilated radioactivity was in fractions other than lipid, protein and free amino acids.) (Data from Table 2 of Febvay et al. (1999)) Incorporation of assimilated sucrose Aphid fraction Respiratory carbon dioxide Lipid Triacylglycerols Phospholipids Protein Free amino acid pool
nmol sucrose mg 1 aphid fresh weight
% of assimilated sucrose
440 285 258 21 138 37
38 24 22 2 12 3
and Crafts-Brandner, 2000) and radiotracer studies confirming that aphids feeding on diets supplemented with 14C-sucrose produce 14C-labelled carbon dioxide (Rhodes et al., 1996; Febvay et al., 1999). Sugars derived from the aphid alimentary tract also enter biosynthetic pathways, leading to the net synthesis of lipid, protein and soluble compounds, including haemolymph sugars and amino acids (Rhodes et al., 1996, 1997; Febvay et al., 1999). Lipids, including phospholipids and especially triacylglycerols, are quantitatively important metabolic products of ingested sucrose. For example, pea aphids feeding on a diet containing 14C-sucrose incorporate 24% of assimilated radioactivity into lipids, double that incorporated into proteins (Table 2). The triacylglycerols of aphids are very unusual in that the chain length of the acyl moieties is small, and dominated by myristic acid (C12) and hexanoic acid (C6) with 1,3-dimyristoyl-2-hexanoyl glycerol as the single most abundant triacylglycerol in A. pisum (Rahbe´ et al., 1994). (The fatty acids in aphid phospholipids are generally longer than in triacylglycerols; for example, the dominant phospholipids of A. pisum are phosphatidylethanolamines and phosphatidylcholines with linoleic acid (C18:2) as their most common acyl moiety (Febvay et al., 1992).) Triacylglycerols serve a crucial role as energy source when aphids are not feeding (and so do not have access to dietary sugars), for example, during moulting and flight, and in the non-feeding adult gynopara and male morphs of certain species. The mobilization of triacylglycerols in aphids has not been studied at the biochemical level, and the processes are presumably comparable to other insects. Briefly, stored triacylglycerols are transformed to diacylglycerols by lipases, as induced by adipokinetic hormones and octopamine, and the diacylglycerols are delivered to different tissues via lipophorin, the principal lipoprotein in insect haemolymph (Canavoso et al., 2001). Consistent with the role of ingested sugars as substrates for lipid synthesis, aphids have no dietary requirement for fatty acids (Dadd, 1985;
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FIG. 12 Fatty acid composition of triacylglycerols in the pea aphid Acyrthosiphon pisum treated with rifampicin to eliminate the symbiotic bacteria (open bars) and untreated aphids (closed bars) reared on chemically defined diets. 14H18: 1-myristoyl-2hexanoyl-3-stearoyl glycerol; 14H16: 1-myristoyl-2-hexanoyl-3-palmitoyl glycerol; 14H14 1,3-dimyristoyl-2-hexanoyl glycerol; 14H12: 1-myristoyl-2-hexanoyl-3-lauroyl glycerol; 14O14 1,3-dimyristoyl-2-octanoyl glycerol; 14S16: 1-myristoyl-2-sorboyl-3palmitoyl glycerol; 14S14: 1,3-dimyristoyl-2-sorboyl glycerol; 14T14: 1,3-dimyristoyl-2octatrienoyl glycerol. (Redrawn from Fig. 3 of Rahbe et al., (1994).)
Douglas, 1988). Furthermore, antibiotic treatment has no effect on the triacylglycerol content and composition of either A. pisum or Macrosiphum euphorbiae (Rahbe´ et al., 1994; Walters et al., 1994) (Fig. 12), indicating that the fatty acids are synthesized by the aphid and not their symbiotic bacteria. The temporal and spatial pattern of sugar metabolism in aphids is largely unknown, and the few datasets available concern the synthesis of haemolymph sugars and proteins. The dominant haemolymph sugar in aphids (as generally for insects) is the disaccharide trehalose, for example at 225 mM in A. pisum (Rhodes et al., 1996). When A. pisum was reared on diets containing 14Csucrose, radioactively labelled trehalose and fructose were recovered from the aphid haemolymph and the ratio of trehalose:fructose increased over the 72 h experiment from 2:3 at 1 h to a stable ratio of 4:1 (Rhodes et al., 1997). In a separate study on A. pisum fed on 14C-sucrose diet over 8 days of larval development, radioactivity was recovered from most or all of the amino acids, depending on the diet composition (Febvay et al., 1999). The synthesis of essential amino acids from ingested sucrose indicates that the assimilated sugar or a metabolic derivative is translocated to the Buchnera cells and used as a substrate for Buchnera-mediated synthesis of essential amino acids, which were then released back to the aphid tissues. Although the hydrolysis products of sucrose are glucose and fructose in equimolar proportions, more fructose than glucose is anticipated to be
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FIG. 13 Mutations in the uricase gene of symbiotic fungi (YLS) of aphids (Tuberaphis species and Glyphinaphis bambusae), as compared with the YLS of the planthopper Nilaparvata lugens. The exons are shown as open boxes, non-sense mutations as asterisks. Tuberaphis species additionally have deletions in the 5’ region, including the TATA box; and T. takenouchii and G. bambusae have frameshift mutations. (From Fig. 6 of Hongoh and Ishikawa (2000).)
assimilated across the gut wall into the aphid tissues because much of the glucose in the ingested sucrose is transformed into oligosaccharides that are voided via the honeydew (Section 4.3.1). A wider array of metabolic paths is available to glucose than fructose, and contributions of fructose to respiration and many biosynthetic pathways, including trehalose synthesis, requires the isomerization of fructose-6-phosphate to glucose-6-phosphate. Two lines of evidence suggest that the metabolic fate of assimilated fructose and glucose may not be equivalent. First, when A. pisum is administered dietary sucrose radioactively labelled in the fructose moiety, less than 10% of the ingested radioactivity is recovered from the aphid tissues and honeydew, raising the possibility that sucrose-derived fructose may be the principal substrate for respiration (Ashford et al., 2000). (This result is in apparent contradiction with the demonstration by Rhodes et al. (1997) of 14C-fructose in the haemolymph of aphids fed on 14C-sucrose, and further study is required to establish whether the discrepancy reflects variation in aphid metabolism or a difference in the sensitivity of different assays.) Second, when Aphis gossypii is reared at high temperature, the polyol mannitol accumulates in the haemolymph; and the conversion of fructose to mannitol by whole insect extracts of A. gossypii suggests that fructose is the precursor of mannitol in vivo (Hendrix and Salvucci, 1998). Galactose is also potentially available to aphids ingesting sugars of the raffinose series, but neither the production of free galactose in the gut lumen of aphids feeding on raffinose sugars nor the assimilation of galactose across the gut wall have been demonstrated.
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Amino acids
Protein synthesis is quantitatively a major fate for amino acids acquired from both the diet and the symbiotic bacteria. For example, in the black bean aphid A. fabae, more than 90% of the total content of most amino acids in aphids is in the protein fraction. Exceptionally, approximately 50% of the aphid tryptophan content is in the free amino acid pool (the reason for this is obscure) (calculated from data in Wilkinson et al., 2001). Some amino acids have important functions other than in protein synthesis. Several amino acids are, or are precursors of, neurotransmitters and neurohormones, e.g. glutamic acid, dopamine (derived from tyrosine), serotonin (derived from tryptophan); and tyrosine is also a major precursor of cuticle synthesis. One consequence of the link between certain amino acids and cell signalling is that the biological ramifications of a shortfall in the supply of these compounds may extend beyond aphid growth and reproductive rates. This issue has not been studied in detail but, as a possible example, 5-hydroxytryptamine, a derivative of dietary tryptophan, has been shown to inhibit the production of alatiform aphids under some experimental conditions (Harrewijn, 1978). Amino acids are also a vital constituent of the haemolymph of aphids. All 20 protein-amino acids are readily detectable in the haemolymph and, as in other insects, the haemolymph amino acids contribute to the osmoregulation of aphid body fluids. Haemolymph amino acids play a crucial role in nitrogen nutrition because the haemolymph is the first destination of both dietary amino acids assimilated across the gut wall and Buchneraderived amino acids released from the mycetocytes. Further research is required to quantify the flux of amino acids into the haemolymph and to establish the specificity and kinetic properties of the amino acid transporters on the gut and mycetocyte membranes. An indication that the flux is high, however, comes from evidence that the haemolymph amino acid pool has very high turnover. In addition to supporting protein synthesis and other functions (see above), haemolymph amino acids are consumed at a high rate in respiration. For example, in plant-reared A. fabae, haemolymph glutamic acid is metabolized to carbon dioxide at a rate of 1.2 nmol carbon mg 1 aphid weight h 1 (Wilkinson et al., 2001) and in diet-reared A. pisum, respiration accounts for 60% of the total glutamic acid assimilated (Febvay et al., 1995). Perhaps amino acids are important to the energy metabolism of particular cell types, even though, in comparison with sucrose, they are minor respiratory substrates for aphids at the level of the whole organism (Rhodes et al., 1996; Febvay et al., 1999). The principal nitrogenous product of amino acid catabolism is ammonia. Aphid honeydew contains appreciable concentrations of ammonia; alternative nitrogenous waste compounds, such as urea and uric acid, are apparently absent (Sasaki et al., 1990; Wilkinson and Douglas, 1995). The symbiotic
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bacteria may contribute to both the elimination of ammonia and the overall nitrogen nutrition of aphids by nitrogen recycling, i.e. the bacteria assimilate aphid waste ammonia into essential amino acids, which are released back to the aphid. Consistent with this proposal, preparations of isolated Buchnera can take up ammonia (Whitehead et al., 1992), and the ammonia content of aphid honeydew is significantly elevated when the symbiotic bacteria are eliminated (Wilkinson and Douglas, 1995). As yet, however, definitive evidence, for example from 15N-tracer experiments, is lacking. The significance of ammonia, despite its toxicity, as the nitrogen excretory product of aphids can be linked to the fact that aphids ingest large volumes of liquid food. Provided the osmotic pressure of this food is reduced soon after ingestion, the conservation of water is not a priority for these insects. The availability of dietary water to aphids may additionally account for absence of Malpighian tubules from aphids and the observation that aphid honeydew and haemolymph are isosmotic, whereas the eliminated material (variously known as urine and faeces) of many terrestrial insects is hyperosmotic relative to the haemolymph. 6.1.3
Sterols
The dominant sterol in aphids, as in other animals, is cholesterol which is both essential to the architecture of cell membranes and precursor of ecdysteroids. Campbell and Nes (1983) have demonstrated that aphid cholesterol is derived from phloem-mobile phytosterols (see Section 2.2.2). As in other insects, the phytosterols are expected to be assimilated from the midgut of aphids and then dealkylated at C24 (Svoboda and Feldlaufer, 1991), probably in the midgut cells prior to distribution to aphid tissues via the haemolymph lipophorin. There is a strong presumption that aphids derive their total sterol requirement from the phloem sap, since neither an aphid nor its complement of symbiotic bacteria can synthesize this class of compounds de novo. Sterol deficiency in insects is generally revealed as dysfunction and death at ecdysis (reflecting a shortfall of ecdysteroids). The fact that these symptoms are not generally observed in aphids, even on unsuitable host plants, suggests that plant phloem sap provides sufficient sterols for aphid nutrition. 6.2 6.2.1
PHYSIOLOGICAL FATE OF ACQUIRED NUTRIENTS
Allocation of nutrients to embryos
Nutrients are allocated preferentially to the reproductive system throughout the life of parthenogenetic aphids. This is a direct consequence of the telescoping of generations in these insects, i.e. embryogenesis is initiated in the larvae and, in some species, even in embryos, such that an adult insect may bear both daughter embryos and, within some of them, grand-daughter
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embryos (Dixon, 1998). As an example, the biomass of embryos in the pea aphid A. pisum increases over larval development by 0.4 g g 1 day 1, twice the rate of the maternal tissues (calculated from data in Douglas, 1996). Nutrients are acquired by the youngest embryos via trophic cords which runs between the embryo and the germarium of the ovary. Older embryos acquire nutrients from the haemolymph (Wilkinson and Ishikawa, 2000). All nutrients required in the diet by the aphid are, by definition, acquired by embryos from the maternal tissues; these nutrients include sterols, various vitamins and minerals as well as sources of organic carbon, nitrogen, phosphorus etc. However, the processes underlying nutrient translocation across the ovarial sheath and into the embryo cells have not been studied. Walters et al. (1994) have suggested that triacylglycerols are a crucial energy source for embryos, but the relative importance of sugars and lipids for energy metabolism of maternal tissues and embryos in aphids has not been investigated systematically. In the reproductive adults of many parthenogenetic aphids, the distribution of embryo size is bimodal: the embryo complement comprises a few large and well-developed embryos and many small, undeveloped embryos (Stadler, 1995). This arrangement can be argued to be of selective advantage to the insect when resources are limiting or unpredictable such that, at any time, nutrients are committed only to those embryos that can be assured of the resources required to develop to parturition. The physiological basis of the controls over the linked processes of the growth/development rate and nutrient uptake by the embryos is unknown, but changes in the metabolic traits of the embryos and transport properties of their epithelial cells (and in the underlying pattern of gene expression) are likely to be important factors mediating the shift from the small to the large embryo class. 6.2.2
The bacterial symbiosis and nutrient allocation patterns
The bacterial symbiosis is both a source and a sink for aphid nutrients, and the flux of metabolites between the Buchnera cells and the mycetocyte (the insect cell containing the Buchnera cells), and between the mycetocytes and the bathing haemolymph, is anticipated to be substantial where aphid protein synthesis rates are high, as in growing larvae and reproductive adults. Various dietary studies have demonstrated that the Buchnera cells provide much or all of an aphid’s entire requirement for many or all of the nine essential amino acids (see Section 5.2.1). The allocation patterns of Buchneraderived essential amino acids are, however, complicated in parthenogenetic aphids by the telescoping of aphid generations (see Section 6.2.1). An adult aphid bears one symbiosis (comprising 60–100 mycetocytes) in her haemocoel, one symbiosis of multiple mycetocytes in each of most of her 50–100 daughter-embryos and, in some aphid species, a symbiosis in one-to-several
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grand-daughter embryos in each of the more mature daughter embryos. Multiple developmentally distinct Buchnera symbioses, therefore, co-exist in a single insect. The simplest route to regulate the nutritional interactions in this complex symbiosis would be for each symbiotic unit to be autonomous, i.e. for the Buchnera cells in each embryo to meet the essential amino acid requirements of that embryo, and the maternal symbiosis to meet the requirements of the maternal tissues, such that there is no net flux of essential amino acids between maternal tissues and embryos. Evidence against this ‘autonomous model’ comes from the demonstration that when the essential amino acids 14C-phenylalanine or 14C-lysine is injected into the haemolymph of A. pisum, radioactivity is recovered from the embryos (Wilkinson and Ishikawa, 1999). These data suggest that the aphid embryos may be a sink for haemolymph phenylalanine and lysine, which are maintained, at least partially, by amino acids synthesized by Buchnera in the maternal tissues, i.e. that the maternal symbiosis subsidizes the essential amino acid nutrition of the embryos. (Definitive demonstration of a maternal subsidy, however, requires checks, first, that the injected essential amino acid has not been metabolized in the maternal tissues prior to uptake of 14C by the embryos and, second, that the incorporation of the amino acids in the embryos reflects net uptake that is not matched by efflux of the same compound by exchange diffusion.) The proposed maternal subsidy may be linked to the lower rates of Buchnera proliferation in final instar larvae and adult aphids than in young larvae and embryos (Whitehead and Douglas, 1993a) for, as nutrients are channelled into Buchnera growth and division, less is available for release to the aphid. This reasoning assumes that the supply of nutrients to Buchnera cells does not vary in parallel with demand. One final point is that, formally, one cannot exclude the possibility that the symbiosis in different mycetocytes may specialize on different essential amino acids, varying with location or developmental age, although there is currently no evidence for such metabolic heterogeneity. Buchnera cells are a metabolic sink for aphid nutrients because they are absolutely dependent on metabolites in the cytoplasm of the mycetocyte for their total nutrient requirements. The requirements of Buchnera are anticipated to include many compounds because of their limited biosynthetic capabilities, including a requirement for phospholipids in membrane synthesis, and all nonessential amino acids for protein synthesis (Shigenobu et al., 2000). Very little is known about nutrient transfer from mycetocyte cytoplasm to Buchnera cells. Experiments on isolated Buchnera preparations suggest that dicarboxylic acids, especially glutamate, are important carbon sources (Whitehead and Douglas, 1993b). Parallel analyses of the free amino acid content of Buchnera cells and mycetocyte cytoplasm revealed that the dominant amino acid in Buchnera is glutamate and in the mycetocytes is -amino butyric acid (GABA) (Whitehead, 1993). Taken together, these data suggest that Buchnera cells derive glutamate from a small, high turnover mycetocyte cytoplasmic pool,
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maintained by the amination of GABA via glutamate dehydrogenase. This metabolic arrangement would combine the dual requirements of supporting the carbon and nitrogen requirements of the Buchnera cells and protecting the aphid nutrient pools from exploitation by Buchnera cells. Other nutrients acquired by Buchnera cells may also be maintained as small, high turnover pools in the mycetocyte cytoplasm. 6.3 6.3.1
DETERMINANTS OF NUTRIENT ALLOCATION PATTERNS
Aphid morph
The nutrient composition of aphids varies with morph. Differences between the alate (winged) and apterous (wingless) morphs of parthenogenetic aphids of the subfamily Aphidinae have been well studied, revealing relatively greater allocation to lipids in the alatiform than apteriform larvae of the final stadium and greater allocation to embryos in adult apterae than in adult alatae (Dixon, 1998). These differences can be interpreted as a trade-off between reproduction and dispersal, reproduction favoured in apterae and dispersal (specifically wing musculature and lipid reserves for flight) favoured in alatae. Aphids are determined as apterae or alatae either as mature embryos or first instar larvae, by crowding and/or nutrient stress, varying among species, but the differences in gene expression and metabolism of key nutrients, e.g. lipid and protein precursors, underlying the distinct developmental pathways remain to be established. Fragmentary information is available on the nutrient allocation patterns in other aphid morphs. The high lipid content of overwintering eggs is indicative of substantial lipid synthesis in sexual females (oviparae). The absence of Buchnera in non-feeding dwarf males and soldiers of various aphid taxa (Buchner, 1965) suggests that the total nutritional requirements, including essential amino acids, in these morphs is met by preformed reserves, but whether nutrient allocation to these morphs differs from feeding morphs bearing Buchnera is unknown. One parthenogenetic generation of various tree-dwelling aphids exhibits distinctive traits of high lipid content, poorly developed gonads, long gut length and low metabolic rate (Dixon, 1973, 1975). This condition is known as reproductive diapause because the aphids produce no offspring for up to several weeks after reaching adulthood, even though they continue to feed and are mobile, and it occurs in the summer months, at a time when the plant phloem sap has particularly high sucrose content and low amino acid content (Wellings and Dixon, 1983; Douglas, 1993). A key feature of reproductive diapause in the sycamore aphid Drepanosiphum platanoidis is the uncoupling of growth of the larva and its complement of embryos, such that the embryo growth and development is arrested at a stage where the head and thorax are incompletely differentiated and the appendages absent (Douglas, 2000).
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The physiological processes underlying the reproductive diapause have not been investigated. 6.3.2
Parasitization
Koinobiont parasitoids dramatically alter the allocation of nutrients in aphids. The eggs and larvae of these insects develop in the body of the living insect and, like a cuckoo in a nest, they divert nutrients from the aphid offspring (embryos) to support their own growth and development. The interaction between A. pisum and the braconid wasp Aphidius ervi has been studied intensively. During oviposition, the female A. ervi injects venom into the aphid, resulting in cessation of aphid oogenesis (i.e. the production of no further aphid embryos), and the growth of pre-existing embryos is also inhibited (Digilio et al., 2000). The bacterial symbiosis is, however, functional, such that the rates of essential amino acid biosynthesis (expressed on a per unit total aphid weight basis) do not differ significantly between parasitized and unparasitized aphids (Rahbe´ et al., 2002). Indeed, parasitization appears to protect the maternal symbiosis from the progressive decline through mycetocyte cell death and depressed growth rates observed in unparasitized aphids, such that the number and total biomass of mycetocytes per aphid is significantly elevated by parasitization for 6–7-day-old aphids (4 days after parasitoid attack) (Cloutier and Douglas, 2003). These data suggest that the parasitoid exploit the nutritional capabilities of the symbiosis to promote its own growth and development. A further indication of the perturbation of nutrient allocation in parasitized aphids is the 3–4 fold increase in the concentration of one amino acid, tyrosine, in the free amino acid pool of A. pisum, at 4–6 days after parasitization by A. ervi (Rahbe´ et al., 2002). Tyrosine is synthesized by the aphid, by decarboxylation of the essential amino acid phenylalanine (including phenylalanine synthesized by Buchnera). The significance of this effect is uncertain. It may arise from parasitoid-mediated manipulation of the profile of amino acids synthesized by the aphid-Buchnera association, linked to a high demand for tyrosine to support cuticle synthesis (Rahbe´ et al., 2002) or, alternatively, low demand for tyrosine in the parasitized aphid resulting in accumulation of this amino acid.
7
Future directions
Recent technical developments have the potential to transform the study of animal nutritional physiology. In particular, it is now possible to identify metabolites and proteins in very small samples and in living cells using advanced analytical methods, e.g. mass spectrometry, nuclear magnetic resonance spectrometry; and molecular/genomic approaches can yield precise
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information on the genetic capabilities of organisms and the impacts of nutritional challenges on gene expression. Research on two specific aspects of the nutritional physiology of aphids and related insects is anticipated to benefit from (and, in some respects, be made possible by) these developments: the regulation of nutrient utilization by aphids (Section 7.1) and the comparative physiology of phloem sap feeding insects (Section 7.2). 7.1
REGULATION OF NUTRIENT UTILIZATION
Insects generally regulate the rate at which they utilize nutrients from their diet by a combination of different processes, including differential digestion and assimilation of dietary constituents, variation in feeding rate and, where different food sources are available, dietary selection (Simpson and Raubenheimer, 1993). Quantitative variation in each of these processes has been demonstrated for a variety of insects, especially orthopterans and lepidopterans, in response to differences in diet composition, nutritional demand, as dictated by developmental age, reproductive status, and environmental circumstance, e.g. temperature (Simpson, 1995) and nonnutritional factors, such as predation risk (e.g. Singer et al., 2002). Some information is available on the contribution of feeding and assimilation rates to the regulation of sugar and amino acid nutrition of aphids on chemically defined diets (Abisgold et al., 1994), but virtually nothing is known about two key aspects of nutritional regulation in aphids: the extent to which food choice at the level of individual sieve elements contributes to the regulation of nutrient input; and how the dietary input of nutrients is integrated with the profile of nutrients acquired from the endogenous bacterial symbionts. These issues are considered in turn, with respect to aphids. 7.1.1
Food choice by aphids
Considered at the level of the whole plant, aphids undoubtedly make food choices. Most aphid species use just one to several related plant genera, and Eastop (1973) concluded that only 10 aphid species can be described as polyphagous; most aphids reject most plant species under most circumstances. Aphids also discriminate across different plant parts, selecting root or shoot, young or mature leaves, flower heads or stems etc. (Dixon, 1998) and they can respond to changing developmental age of plant parts by ‘fine-scale migration’ between different plant parts (Harrington and Taylor, 1990). An important unresolved question is whether aphids also discriminate among the various sieve elements at their site of feeding. It is evident from many EPG datasets on various aphid/plant species combinations that aphids do not necessarily feed from the first sieve element penetrated by their stylets. Several explanations can be put forward to explain the withdrawal of the stylets shortly after penetrating a sieve element. Perhaps
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the aphid is responding to the chemical composition of the phloem sap, including the concentration of key nutrients as detected by the epipharyngeal gustatory organ (see Fig. 5b), and/or the hydrostatic pressure of the sieve element contents, which would drop precipitately if the sieve element sealed in response to aphid puncture. Multiple penetrations and the associated release of saliva into the sieve element may also suppress the capacity of the plant vascular tissue generally or the individual sieve element punctured to respond to stylet puncture; this hypothesized function of multiple penetrations has been described as ‘conditioning’, but the underlying molecular and biochemical processes are unknown (see Section 3.3). Resolution among these alternatives would contribute to our understanding of whether aphids discriminate among sieve elements. More information on the composition of phloem sap would establish whether it is advantageous for aphids to select particular sieve elements. For example, the diurnal variation in phloem sugars and amino acids is not trivial (Section 2.2). If the diurnal variation in sap composition in any one sieve element were considerably greater than the among-element variation at any one time, then it may not be advantageous for an aphid to discriminate between sieve elements on the basis of nutrient levels. An indication that aphids may not discriminate comes from the finding that the amino acid concentration in phloem exudates from severed aphid stylets on pea and wheat can vary several-fold (see Section 2.2.3); if aphids selected sieve elements on the criterion of amino acid concentration, then the exudate concentration would not be expected to display so great a variation. Also, the close similarity between amino acid composition of EDTA exudates and severed stylet exudates of phloem sap (Weibull et al., 1990) suggests that the sieve elements selected by aphids are not nutritionally atypical, at least with respect to amino acids. Establishing the extent, first, of inter-sieve element variation in nutrient composition and, second, to which aphids discriminate among sieve elements on nutritional criteria would contribute to our understanding of the importance of post-ingestive factors in the match between dietary supply of nutrients and aphid demand for growth and reproduction. The increasing precision of analyses of sieve element physiology, including the use of confocal microscopy to study processes in vivo (e.g. Van Bel et al., 2002), and (as mentioned in the opening paragraph of Section 7) the greatly enhanced sensitivity of recent methods to quantify plant metabolites in very small volumes (e.g. Tomos and Sharrock, 2001) provide the technical bases to resolve these issues. 7.1.2
Integration of the bacterial symbiosis into the nutritional physiology of aphids
Essential amino acids are crucial to the nitrogen nutrition of aphids. If one essential amino acid is in short supply, protein synthesis by the aphid is
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curtailed, resulting in depressed growth and reproduction, whatever the availability of the other eight essential amino acids. Aphids derive much of their essential amino acid requirement from Buchnera cells but, as considered in Section 5.2.2, sequence data for Buchnera suggest that production of most essential amino acids is constitutive, i.e. the rate of synthesis is not responsive to concentration of the product. The implication is that the essential amino acid nutrition of aphids depends on aphid-mediated integration of a fixed supply from Buchnera and variable dietary supply. In particular, the regulation of the rate of amino acid assimilation from the gut (Section 4.4.1) and the utilization of amino acids as respiratory substrates (Section 6.1.2) could be crucial, but as yet barely explored, determinants of the protein synthesis rates of aphids. This scenario, however, pre-supposes that the most parsimonious interpretation of Buchnera sequence data is correct. However, sequence data alone are inadequate to infer function accurately. A recent study of the trpEG genes of Buchnera provides a cautionary tale. In many aphids, including the Aphidinae, trpEG genes are amplified relative to other genes coding for enzymes in the tryptophan biosynthetic pathway (Lai et al., 1994; Van Ham et al., 1999), for example 2–8 fold in A. pisum, varying between clones (Birkle et al., 2002). TrpEG code for anthranilate synthase, the enzyme traditionally regarded as the pacemaker enzyme in tryptophan synthesis, i.e. its activity dictates the rate of tryptophan synthesis, generating the expectation that tryptophan synthesis rates increase with increasing trpEG copy number. This expectation was, however, not upheld by direct quantification of tryptophan synthesis, which varied significantly among aphid lines but independently of their trpEG copy number (Birkle et al., 2002). Should the processes controlling essential amino acid synthesis by Buchnera be evident in the Buchnera sequence? If, as suggested in Section 6.1.2, essential amino acid synthesis is substrate-limited, i.e. regulated by the supply of precursors, possibly as determined by transporter function on the aphid membrane bounding each Buchnera cell, then the predicted genomic evidence would be negative; the decay and loss of bacterial regulatory sequences present in free-living relatives of Buchnera, such as E. coli. The sequence data are consistent with this expectation. However, information on the extent and mode of regulation of essential amino acid synthesis by Buchnera requires combined molecular and metabolic approaches, including consideration of the nutritional physiology of the insect partner. The general conclusion is two-fold: the need for great care in interpreting putative function from genomic data for Buchnera; and need to address the utilization of dietary amino acids and symbiosis-derived amino acids as elements in a single study and not as separate topics. The possibility that the regulatory networks determining the rate of Buchnera-mediated essential amino acid synthesis may be located exclusively to the aphid compartment is reminiscent of the condition of bacterial-derived
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organelles (mitochondria and plastids). Is Buchnera an essential amino acid synthesizing organelle? Mitochondria and plastids have much reduced genomes relative to their likely free-living ancestors and can be defined as organelles because genes essential to their function have, in the course of evolution, been transferred to the host nucleus, such that they are dependent on the nucleocytoplasm for sustained function (Douglas and Raven, 2003). The Buchnera genome at 0.45–0.64 Mbp (Shigenobu et al., 2000; Wernegreen et al., 2001; Gil et al., 2002) is intermediate in size between free-living bacteria, such as E. coli at 4.95 0.25 Mbp (Bergthorsson and Ochman, 1998) and the plastids (0.12–0.20 Mbp)/mitochondria (generally 0.014–0.070 Mbp, but up to 2 Mb in plants); and it is not known whether the aphid genome contains Buchnera-derived genes whose products are targeted back to the Buchnera cells and are essential to Buchnera function. However, the possible transfer of the genes ilvA and metC to the aphid nucleus is one among several explanations for the capacity of Buchnera to synthesize isoleucine and methionine, respectively, despite the absence of sequence in the Buchnera genome attributable to these genes (see Section 5.2.1). Understanding the evolutionary status of Buchnera as a bacterium, organelle or intermediate between these two conditions will contribute to a proper grasp of the processes by which symbiotic amino acid production is integrated into the overall nitrogen nutrition of aphids. Mitochondria and plastids have acquired functions apparently absent from their bacterial ancestors, such as fatty acid metabolism, calcium storage and a role in apoptosis, presumably linked to their declining control over their own metabolic pools and signalling networks. These organelles are membranebound compartments in eukaryotic cells to which specific functions have been allocated. Do Buchnera cells provide analogous service, potentially important to the nutritional physiology of the aphid? In this respect, some attributes of aphids experimentally deprived of their complement of Buchnera (known as aposymbiotic aphids) deserve careful consideration. For example, does the depressed uptake of the essential amino acid leucine across the gut wall of aposymbiotic aphids (Douglas et al., 2001) reflect a direct involvement of Buchnera in amino acid assimilation or a secondary consequence of depressed protein synthesis (with the implication that the gut amino acid transporters of aphids are proteins with very high turnover)? Such issues can only be resolved by concerted analysis of the nature and scale of integration of the Buchnera cells into the wider nutritional physiology of the aphid. 7.2 7.2.1
COMPARATIVE PHYSIOLOGY OF PHLOEM SAP FEEDERS
The diversity of phloem sap-feeding animals
Phloem sap is a ‘difficult’ food source. An animal specialized for phloem sap feeding must, first, be small enough to tap into intact sieve elements without
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disturbing the sap stream and, second, display anatomical and physiological adaptations to the nutritionally unbalanced composition and high osmotic and hydrostatic pressure of phloem sap. The Hemiptera is apparently the sole group of animals with members that utilize plant phloem sap as their principal or sole source of food through their life cycle. The phloem-feeding habit has evolved independently multiple times in this group, at least twice in the Heteroptera and at least once in the ancestor of each of the Auchenorrhyncha and Sternorrhyncha (see legend to Fig. 1 for information on the taxonomy of the Hemiptera). Other animals that utilize phloem sap include thrips, lepidopterans, hummingbirds and primates (including those humans who like maple syrup) (e.g. Daily et al., 1993; Passamani and Rylands, 2000) but, for all of these animals, phloem sap is a quantitatively small and non-required element of their diet. What features of the Hemiptera have led to their remarkable pre-disposition for the phloem sap feeding habit? Two important characteristics are the anatomical specialization of their mouthparts for feeding on liquid food (Dolling, 1991) and the apparent facility with which these insects form intimate symbioses with micro-organisms which provide nutrients (Douglas, 1989). Neither of these traits would, however, preclude the evolution of phloem sap feeding in other insect groups. Mouthparts well-suited for imbibing liquid food have evolved in various non-hemipteran insects, including members of the Diptera, Hymenoptera and Lepidoptera. Many insects bear symbiotic microorganisms (Douglas, 1989); most notably the blood-feeding dipterans Glossina spp. (tse tse flies) whose symbionts, Wigglesworthia sp., are very closely related to Buchnera, the symbionts of aphids (Chen et al., 1999). The anatomy of the alimentary tract has been proposed by Goodchild (1966) as a factor contributing to the restriction of phloem sap feeding to Hemiptera. The Hemiptera lacks a crop, i.e. dilated portion of the foregut, in which ingested phloem sap could accumulate, causing osmotic stress (specifically, loss of body water to the crop as a consequence of the high osmotic pressure of phloem sap). Goodchild (1966) considers that the possession of a crop (as, for example, in the Diptera) would ‘render the occasional piercing of phloem bundles more frequently fatal to the insect’. This perspective may, however, be an oversimplification, as it is well-established that various non-hemipteran insects can utilize foods of extreme osmotic pressure or water content (Edney, 1977). Furthermore, the remarkable diversity of gut anatomy among the Hemiptera including phloem sap feeding forms, all interpreted to function in controlling water flux between the gut lumen and body fluids (Goodchild, 1966), suggests that at least some of the anatomical ‘problems’ associated with phloem sap feeding can be solved in multiple different ways. This topic is ready for re-examination because the array of methodologies available to explore the factors determining the narrow phylogenetic distribution of specialized phloem sap feeding is far greater today than when Goodchild (1966) addressed this issue. In particular, the nutritional physiology
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of small insects has been transformed by techniques now available to quantify metabolites in nano- to pico-volumes and molecular approaches (see Section 7.2.2); and sophisticated phylogenetic methods can be applied to explore the evolutionary origins of complex traits. 7.2.2
Molecular physiology of the symbiosis between phloem-feeding insects and micro-organisms
Molecular approaches can both promote understanding of the mechanisms underlying physiological processes and make it possible to discriminate between evolutionarily conserved and convergent processes across different taxa of insects feeding on phloem sap. Access to the complete genome sequence of the organism under study transforms the physiological opportunities available, but much can be achieved while awaiting the complete sequence of an aphid. To date, very few nutritional physiological studies of phloem-feeding Hemiptera have exploited molecular techniques, but the potential of molecular biology in the comparative physiological approach is illustrated by two investigations, described below. The first study concerns the accumulation of polyols in insects exposed to elevated temperatures: mannitol in the aphid Aphis gossypii and sorbitol in the whitefly B. argentifolii (Hendrix and Salvucci, 1998). Sorbitol synthesis in B. argentifolii is mediated by a biochemically unusual NADPH-ketose reductase/ sorbitol dehydrogenase (Wolfe et al., 1998), the cDNA of which has been identified and sequenced (Wolfe et al., 1999). The production of mannitol by temperature-stressed A. gossypii suggests that the ketose reductase/polyol dehydrogenase may not be unique to whitefly; and an understanding of the phylogenetic distribution of this response to high temperature could be obtained by investigating the enzymological and genetic basis of polyol synthesis among hemipterans and the impact of temperature on the expression patterns of the relevant gene(s). The research on sorbitol production in B. argentifolii is exceptional for phloem sap feeding insects in that data on all of the physiology, biochemistry and sequence are available (albeit for just one insect species). Although there is no indication that the capacity for polyol accumulation in the haemolymph is restricted to insects feeding on phloem sap, the elevated sorbitol concentrations in the haemolymph of B. argentifolii reared on high sucrose diets at standard temperature (Wolfe et al., 1998) suggest that polyol production may contribute to osmoregulation, a crucial attribute for phloem sap feeders because of the high and variable osmotic pressure of phloem sap. One important example of physiological variation among phloem sap feeding Hemiptera is the principal nitrogenous excretory compound, which is, for example ammonia in aphids (see Section 6.1.2) and uric acid in planthoppers. Planthoppers of the family Delphacidae bear symbiotic fungi,
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informally known as yeast-like symbionts (YLS). The YLS have functional uricase activity and contribute to the nitrogen economy of the insect by utilizing uric acid (Sasaki et al., 1996). Members of this clade of fungi are also found in the aphid tribe Cerataphini (which have secondarily lost Buchnera, the usual symbiont of aphids). The principal nitrogenous waste product of the Cerataphini is presumably ammonia, as in other aphids, and the selection pressure for functional uricase of their YLS is much reduced and possibly absent. Consistent with this expectation, the uricase gene in the YLS of aphids bear a frame-shift or non-sense mutations and large deletions in the 5’-flanking region (Fig. 13), all indicative of loss of function (Hongoh and Ishikawa, 2000). There is, thus, an excellent correspondence between gene sequence of the symbiont and insect physiology, with respect to uricase. Sequence data alone should be used to infer function with caution, as considered in Section 7.1.2. Even so, the study of Hongoh and Ishikawa (2000) illustrates how the study of single genes of one clade of symbiotic microorganisms in different insect hosts can contribute to our understanding of the physiological similarities and differences among phloem-feeding insects. The comparative genomic analysis of Buchnera in two aphid species (Shigenobu et al., 2000; Tamas et al., 2002) offers the first extension of this approach to complete genomes. In this way, symbiosis, which is integral to the nutritional physiology of phloem-feeding insects, can also become integral to the study of their nutritional physiology.
8
Concluding comments: why study the nutritional physiology of phloem sap feeding insects
Why study the nutritional physiology of phloem sap feeding insects? There are two reasons beyond ‘that they exist’. The first is that phloem sap is an extreme diet, as defined by its utilization as principal or sole source of food by a very low diversity of animals, apparently only hemipteran insects. We have some insight into the barriers to phloem sap utilization overcome by these insects; they include nutritional imbalance, overcome at least partly by acquisition of a microbial symbiosis, and high osmotic pressure overcome by anatomical modifications, gut transglucosidase activity and probably ion transport functions. Importantly, however, phloem sap differs from most extreme diets in that it is part of living organisms, which can respond through natural selection to the negative consequences of insect consumption. (Vertebrate blood has parallels with plant phloem sap in this respect, although it is less ‘extreme’ than phloem sap in that it is utilized by a diversity of animals, including members of several insect orders, e.g. Diptera, Siphonaptera and Hemiptera.) The significance of animals, especially hemipterans, as a selection pressure moulding the composition of phloem sap (especially its high ratios of
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sugars:amino acids and non-essential:essential amino acids) and sieve element organization is, at present, an open question. Equally, the relative contribution of the phloem sap and other plant features (especially surface characteristics and interactions with natural enemies) to the ongoing antagonistic coevolution between plants and phloem sap feeding insects is unclear. Although much of current research on plant resistance to phloem sap feeders concerns plant features unrelated to phloem sap utilization and insect nutritional physiology, it is uncertain whether this reflects fundamental constraints on the evolutionary diversification in sieve elements and phloem sap related to the requirements for sustained, long-distance transport or the greater ease with which interactions at the plant surface can be studied relative to interactions at the level of the sieve element. These issues are crucial to addressing the contribution of nutritional physiology to the related discipline of nutritional ecology of phloem sap feeding insects, including the extent to nutritional factors shape plant range, life history traits etc. An added impetus to this topic is provided by the increasing evidence for large impacts of climate change on populations of phloem sap feeders (e.g. Percy et al., 2002); the physiological processes underlying the projected shifts in insect–plant interactions are only weakly understood. A second reason to study the nutritional physiology of phloem sap feeding insects is that this discipline encompasses traits potentially suitable as targets in novel pest management strategies specific to these insects, i.e. traits linked to their feeding habits. At the whole-insect level, potential targets can be identified readily: the gut sucrase/transglucosidase activity of aphids on which the insect carbon nutrition and osmoregulation depend, the proliferation of and nutrient release from symbiotic micro-organisms, and possibly transporter function for assimilation of nutrients from the gut lumen. At the molecular and biochemical levels, all these processes are unknown but are resolvable in the context of recent advances in analytical and genomic methodologies (see Section 7). The pest control opportunities afforded by nutritional physiology are, by definition, related to nutrition and dependent on insect feeding. These approaches to pest management will depress pest populations and are not anticipated to achieve ‘knock-down’, as obtained with conventional broadspectrum insecticides with the nervous system as their primary target. They have greatest potential as part of an integrated pest management strategy; and the specificity of the anticipated strategies to phloem sap feeders will be of particular value in this context. The most likely mode of delivery of active agents is the oral route via phloem sap, for which GM technology of crop plants would be well suited, but not necessarily required. The immediate outlook for novel pest management strategies exploiting the unique features of phloem sap feeding insects is poor, with declining commercial research effort in crop plant protection and an antipathy to GM, at least in Europe. This should not deter the fundamental research on the
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nutritional physiology of phloem sap feeders that must precede application in the agricultural context. Novel pest management strategies will be needed to combat the increasing incidence of resistance to conventional insecticides among aphids and whitefly, to meet the increasing demand for more specific insecticides for environmental and public health reasons, and to support the anticipated increased crop production required to feed the projected human population of nine billion. Realization of these novel pest management strategies depends on sustained research on the nutritional physiology of phloem sap feeding pests.
References Abisgold, J. D., Simpson, S. J. and Douglas, A. E. (1994). Nutrient regulation in the pea aphid Acyrthosiphon pisum – application of a novel geometric framework to sugar and amino acid consumption. Physiol. Ent. 19, 95–102. Antognoni, F., Fornale, S., Grimmer, C., Komor, E. and Bagni, N. (1998). Longdistance translocation of polyamines in phloem and xylem of Ricinus communis L. plants. Planta 204, 520–527. Ashford, D. A., Smith, W. A. and Douglas, A. E. (2000). Living on a high sugar diet: the fate of sucrose ingested by a phloem-feeding insect, the pea aphid Acyrthosiphon pisum. J. Insect Physiol. 46, 335–342. Auclair, J. L. (1963). Aphid feeding and nutrition. Annu. Rev. Entomol. 8, 439–490. Baumann, P., Baumann, L. and Clark, M. A. (1994). Levels of Buchnera aphidicola chaperonin GroEL during growth of the aphid Schizaphis graminum. Curr. Microbiol. 32, 279–285. Bergthorsson, U. and Ochman, H. (1998). Distribution of chromosome length variation in natural isolates of Escherichia coli. Mol. Biol. Evol. 15, 6–16. Berlandier, F. A. (1996). Alkaloid levels in narrow-leafed lupin, Lupinus angustifolius, influences green peach aphid reproductive performance. Entomol. Exp. Appl. 79, 19–24. Birkle, L. M., Minto, L. B. and Douglas, A. E. (2002). Relating genotype and phenotype for tryptophan synthesis in an aphid-bacterial symbiosis. Physiol. Ent. 27, 302–306. Blackman, R. L. and Eastop, V. F. (1984). Aphids on World Crops, pp. 465. Chichester, UK: John Wiley. Botha, C. E. J, Malcolm, S. B. and Evert, R. F. (1977). An investigation of preferential feeding habit in four Asclepiadaceae by the aphid, Aphis nerii B. de F. Protoplasma 92, 1–19. Bourgis, F., Rije, S., Nuccio, M. L., Fisher, D. B., Tarczynski, M. C., Li, C., Herschbach, C., Rennnenberg, H., Pimenta, M. J., Shen, T-L., Gage, D. A., Hanson, A. D. (1999). S-methylmethionine plays a major role in phloem sulphur transport and is synthesized by a novel type of methyltransferase. Plant Cell 11, 1485–1497. Bridges, M., Jones, A. M. E., Bones, A. M., Hodgson, C., Cole, R., Bartlet, E., Wallsgrove, R., Karapapa, V. K., Watts, N., Rossiter, J. T. (2002). Spatial organisation of the glucosinolate-myrosinase system in brassica specialist aphids is similar to that of the host plant. Proc. Roy. Soc. Lond. B 269, 187–191. Brough, C. N. and Dixon, A. F. G. (1990). Ultrastructural features of egg development in oviparae of the vetch aphid, Megoura viciae Buckton. Tissue Cell 22, 51–63.
THE NUTRITIONAL PHYSIOLOGY OF APHIDS
131
Brudenell, A. J. P., Griffiths, H., Rossiter, J. T. and Baker, D. A. (1999). The phloem mobility of glucosinolates. J. exp. Bot. 50, 745–756. Brunings, E. A. and DePriester, W. (1971). Effects of mode of fixation on formation of extrusions in the midgut epithelium of Calliphora erythrocephala (Diptera: Calliphoridae). Cytobiologie 4, 487–492. Buchner, P. (1965). Endosymbiosis of Animals with Plant Micro-organisms, pp. 909. Chichester: Wiley. Caillaud, C. M. and Niemeyer, H. M. (1996). Possible involvement of the phloem sealing system in the acceptance of a plant as host by an aphid. Experientia 52, 927–931. Campbell, B. C. and Nes, W. D. (1983). A reappraisal of sterol biosynthesis and metabolism in aphids. J. Insect Physiol. 29, 149–156. Campbell, B. C., Steffen-Campbell, J. D., Sorensen, J. T. and Gill, R. J. (1995). Paraphyly of Homoptera and Auchenorrhyncha inferred from 18S rDNA nucleotide sequences. Syst. Entomol. 20, 175–194. Canavoso, L. E., Jouni, Z. E., Karnas, K. J., Pennington, J. E. and Wells, M. A. (2001). Fat metabolism in insects. Annu. Rev. Nut. 21, 23–46. Castagna, M., Shayakul, C., Trotti, D., Sacchi, V. F., Harvey, W. R. and Hediger, M. A. (1998). Cloning and characterization of a potassium-coupled amino acid transporter. Proc. Natl Acad. Sci. USA 95, 5395. Chen, D. Q., Montllor, C. B. and Purcell, A. H. (2000). Fitness effects of two facultative endosymbiotic bacteria on the pea aphid, Acyrthosiphon pisum, and the blue alfalfa aphid, A. kondoi. Entomol. Exp. Appl. 95, 315–323. Chen, S., Petersen, B. L., Olsen, C. E., Schulz, A. and Halkier, B. A. (2001). Longdistance phloem transport of glucosinolates in Arabidopsis. Plant Physiol. 127, 194–201. Chen, X. A., Li, S. and Aksoys, S. (1999). Concordant evolution of a symbiont with its host insect species: Molecular phylogeny of genus Glossina and its bacteriomeassociated endosymbiont, Wigglesworthia glossinidia. J. mol. Evol. 48, 49–58. Cloutier, C. and Douglas, A. E. (2003). Impact of a parasitoid on the bacterial symbiosis of its aphid host. Entomol. Exp. Appl. (in press). Corbesier, L., Havelange, A., Lejeune, P., Bernier, G. and Pe´rilleux, C. (2001). N content of phloem and xylem exudates during the transition to flowering in Sinapis alba and Arabidopsis thaliana. Plant, Cell Env. 24, 367–375. Cristofoletti, P. T., Ribeiro, A. F., Deraison, C., Rahbe, Y. and Terra, W. R. (2003). Midgut adaptation and digestive enzyme distribution in a phloem feeding insect, the pea aphid Acyrthosiphon pisum. J. Insect Physiol. 49, 11–24. Dadd, R. H. (1985). Comprehensive insect physiology, biochemistry and pharmacology. In: Nutrition: organisms (eds. Kerkut, G. A., and Gilbert, L. I.) Vol. 4. pp. 313–391. Oxford: Pergamon. Daily, G. C., Ahrlich, P. R. and Haddad, N. M. (1993). Double keystone bird in a keystone species complex. Proc. Natl. Acad. Sci. USA 90, 592–594. Darby, A. C., Birkle, L. M., Turner, S. L. and Douglas, A. E. (2001). An aphid-borne bacterium allied to the secondary symbionts of whitefly. FEMS Microbiol. Ecol. 36, 43–50. Digilio, M. C., Isidoro, N., Tremblay, E. and Pennacchio, F. (2000). Host regulation effects of ovary fluid and venom of Aphidius ervi Haliday (Hymenoptera: Braconidae). J. Insect Physiol. 46, 1041–1050. Dixon, A. F. G. (1973). Metabolic acclimatization to seasonal changes in temperature in the sycamore aphid, Drepanosiphum platanoides (Schr.) and lime aphid, Eucallipterus tiliae L. Oecologia 13, 205–210.
132
A. E. DOUGLAS
Dixon, A. F. G. (1975). Seasonal changes in fat content, form, state of gonads, length of adult life in the sycamore aphid Drepanosiphum platanoides (Schr.). Trans. Roy. Entomol. Soc. Lond. 127, 87–99. Dixon, A. F. G. (1998). Aphid Ecology, pp. 300. Chapman and Hall, London. Dolling, W. R. (1991). The Hemiptera, pp. 274. Oxford, UK: Oxford University Press. Douglas, A. E. (1988). Sulphate utilisation in an aphid symbiosis. Insect Biochem. 18, 599–605. Douglas, A. E. (1989). Mycetocyte symbiosis in insects. Biol. Revs 69, 409–434. Douglas, A. E. (1990). Nutritional interactions between Myzus persicae and its symbionts. In: Aphid-Plant Genotype Interactions (eds. Campbell, R. K., and Eikenbary, R. D.), pp. 319–327. Amsterdam: Elsevier. Douglas, A. E. (1992). Requirement of pea aphids (Acyrthosiphon pisum) for their symbiotic bacteria. Entomol. Exp. Appl. 65, 195–198. Douglas, A. E. (1993). The nutritional quality of phloem sap utilized by natural aphid populations. Ecol. Entomol. 18, 31–38. Douglas, A. E. (1996). Reproductive failure and the free amino acid pools in pea aphids (Acyrthosiphon pisum) lacking symbiotic bacteria. J. Insect Physiol. 42, 247–255. Douglas, A. E. (2000). Reproductive diapause and the bacterial symbiosis in the sycamore aphid Drepanosiphum platanoidis. Ecol. Entomol. 25, 256–261. Douglas, A. E. and Prosser, W. A. (1992). Synthesis of the essential amino acid tryptophan in the pea aphid (Acyrthosiphon pisum) symbiosis. J. Insect Physiol. 38, 565–568. Douglas, A. E. and Raven, J. A. (2003). Genomes at the interface between bacteria and organelles. Phil. Trans. Roy. Soc. Lond. B 358, 5–17. Douglas, A. E., Minto, L. B. and Wilkinson, T. L. (2001). Quantifying nutrient production by the microbial symbiosis in an aphid. J. Exp. Biol. 204, 349–358. Douglas, A. E., Darby, A. C., Birkle, L. M. and Walters, K. F. A. (2002). The ecological significance of symbiotic micro-organisms in animals: perspectives from the microbiota of aphids. In: Genes in the Environment (eds. Hails, R. M., Beringer, J., and Godfray, H. C. J.), pp. 306–325. Oxford: Blackwell Scientific Publishers. Down, R. E., Ford, L., Woodhouse, S. D., Raemaekers, R. J. M., Leitch, B., Gatehouse, J. A. and Gatehouse, A. M. R. (2000). Snowdrop lectin (GNA) has no acute toxic effects on a beneficial insect predator, the 2-spot ladybird (Adalia bipunctata L.). J. Insect Physiol. 46, 379–391. Duspiva, F. (1955). Enzymatische Prozesse bei der honingtaubuldung der Aphiden. Vergl. Dtsch. Zool. Ges. 18, 440–447. Eastop, V. F. (1973). Diversity of the Sternorrhyncha within major climatic zones. In: Diversity of Insect Faunas (eds. Mound, L. A., and Waloff, N.), pp. 71–88. Oxford: Blackwell Scientific Publications. Edney, E. B. (1977). Water Balance in Land Arthropods, pp. 156. Berlin: Springer Verlag. Edwards, J. S. (1966). On the nature and origin of solid components in the gut contents of aphids. J. Insect Physiol. 12, 31–35. Febvay, G., Pageaux, J. F. and Bonnot, G. (1992). Lipid composition of the pea aphid, Acyrthosiphon pisum (Harris) (Homoptera, Aphididae), reared on host plant and on artificial media. Arch. Insect Biochem. 21, 103–118. Febvay, G., Liadouze, I., Guillaud, J. and Bonnot, G. (1995). Analysis of energetic amino acid metabolism in Acyrthosiphon pisum: a multidimensional approach to amino acid metabolism in aphids. Arch. Insect Biochem. Physiol. 29, 45–69. Febvay, G., Rahbe, Y., Rynkiewiecz, M., Guillaud, J. and Bonnot, G. (1999). Fate of dietary sucrose and neosynthesis of amino acids in the pea aphid, Acyrthosiphon pisum, reared on different diets. J. Exp. Biol. 202, 2639–2652.
THE NUTRITIONAL PHYSIOLOGY OF APHIDS
133
Fisher, D. B. (2000). Long-distance transport. In: Biochemistry and Molecular Biology of Plants (eds. Buchanan, B., Gruissem, W., and Jones, R.), pp. 730–784. Rockville, MD, USA: American Society of Plant Physiologists. Foissac, X., Edwards, M. G., Du, J. P., Gatehouse, A. M. R. and Gatehouse, J. A. (2002). Putative protein digestion in a sap-sucking homopteran plant pest (rice brown plant hopper; Nilaparvata lugens: Delphacidae) – identification of trypsinlike and cathepsin B-like proteases. Insect Biochem. Mol. Biol. 32, 967–978. Forrest, J. M. S. (1987). Galling aphids. In: Aphids Their Biology, Natural Enemies and Control (eds. Minks, A. K., and Harrewijn, P.) Vol. 1. pp. 341–354. Amsterdam: Elsevier. Forrest, J. M. S. and Knight, B. A. (1972). Presence of phytosterols in the food of the aphid, Myzus persicae. J. Insect Physiol. 18, 723–728. Fritz, E., Evert, R. F. and Heyser, W. (1983). Microautoradiographic studies of phloem loading and transport in the leaf of Zea mays L. Planta 159, 193–206. Fukatsu, T., Nikoh, N., Kawai, R. and Koga, R. (2000). The secondary endosymbiotic bacterium of the pea aphid Acyrthosiphon pisum (Insecta: Homoptera). Appl. Env. Microbiol. 66, 2748–2758. Gange, A. C. (1995). Aphid performance in an alder (Alnus) hybrid zone. Ecology 76, 2074–2083. Gatehouse, J. A. (2002). Plant resistance towards insect herbivores: a dynamic interaction. New Phytol. 156, 145–169. Genger, D. R. and Servaites, J. C. (1994). Diurnal regulation of photosynthetic carbon metabolism in C3 plants. Annu. Rev. Pl. Physiol. Mol. Biol. 45, 235–256. Gil, R., Sabater-Munoz, B., Latorre, A., Silva, F. J. and Moya, A. (2002). Extreme genome reduction in Buchnera spp.; toward the minimal genome needed for symbiotic life. Proc. Natl. Acad. Sci. USA 99, 4454–4458. Girousse, C., Bournoville, R. and Bonnemain, J-L. (1996). Water deficit-induced changes in concentrations in proline and some other amino acids in the phloem sap of alfalfa. Plant Physiol. 111, 109–113. Givovich, A., Sandstro¨m, J., Niemeyer, H. M. and Petterson, J. (1994). Presence of a hydroxamic acid glucoside in wheat phloem sap, and its consequences for the performance of Rhopalosiphum padi (L.) (Homoptera: Aphididae). J. Chem. Ecol. 20, 1923–1930. Goodchild, A. J. P. (1966). Evolution of the alimentary canal in the Hemiptera. Biol. Revs. 41, 97–140. Grenier, A-M., Nardon, C. and Rahbe, Y. (1994). Observations on the microorganisms occurring in the gut of the pea aphid Acyrthosiphon pisum. Entomol. Exp. Appl. 70, 91–96. Ha¨flinger, B., Kindhauser, E. and Keller, F. (1999). Metabolism of D-glycero-Dmanno-heptitol, volemitol, in polyanthus. Discovery of a novel ketose reductase. Plant Physiol. 119, 191–197. Harada, H. and Ishikawa, H. (1997). Experimental pathogenicity of Erwinia aphidicola to pea aphid, Acyrthosiphon pisum. J. Gen. Appl. Microbiol. 43, 363–367. Harada, H., Oyaizu, H. and Ishikawa, H. (1996). A consideration about the origin of aphid intracellular symbiont in connection with gut bacterial flora. J. Gen. Appl. Microbiol. 42, 17–26. Harrewijn, P. (1978). The role of plant substances in polymorphism of the aphid Myzus persicae. Entomol. Exp. Appl. 24, 198–214. Harrington, R. and Taylor, L. R. (1990). Migration for survival: fine scale population redistribution in an aphid Myzus persicae. J. Anim. Ecol. 59, 1177–1193. Hartmann, T. (1999). Chemical ecology of pyrrolizidine alkaloids. Planta 207, 483–495.
134
A. E. DOUGLAS
Hartmann, T. and Dierich, B. (1998). Chemical diversity and variation of pyrrolizidine alkaloids of the senecionine type: biological need or co-incidence?. Planta 206, 443–451. Hawthorne, D. J. and Via, S. (2001). Genetic linkage of ecological specialization and reproductive isolation in pea aphids. Nature 412, 904–907. Hayashi, H., Fukuda, A., Suzui, N. and Fujimaki, S. (2000). Proteins in the sieve element-companion cell complexes: their detection, localisation and possible functions. Aust. J. Plant Physiol. 27, 489–496. Hendrix, D. L. and Salvucci, M. E. (1998). Polyol metabolism in homopterans at high temperatures: accumulation of mannitol in aphids (Aphididae: Homoptera) and sorbitol in whiteflies (Aleyrodidae: Homoptera). Comp. Biochem. Physiol. 20A, 487–494. Hendrix, D. L., Wei, Y-A. and Leggett, J. E. (1992). Homopteran honeydew sugar composition is determined by both the insect and plant species. Comp. Biochem. Physiol. 101B, 23–27. Hinde, R. (1971). The control of the mycetome symbiotes of the aphid Brevicoryne brassicae, Myzus persicae and Macrosiphum rosae. J. Insect Physiol. 17, 1791–1800. Hongoh, Y. and Ishikawa, H. (2000). Evolutionary studies on uricases of fungal endosymbionts of aphids and planthoppers. J. Mol. Evol. 51, 265–277. House, H. L. (1974). Digestion. In: The Physiology of Insecta, 2nd edition (Rockstein, M.), pp. 63–117. New York: Academic Press. Hu, H., Penn, S. G., Lebrilla, C. B. and Brown, P. H. (1997Isolation and characterization of soluble B-complexes in higher plants. The mechanism of phloem mobility of boron. Plant Physiol. 113, 649–655. Humphreys, N. J. and Douglas, A. E. (1997). The partitioning of symbiotic bacteria between generations of an insect: a quantitative study of Buchnera in the pea aphid (Acyrthosiphon pisum) reared at different temperatures. Appl. Env. Microbiol. 63, 3294–3296. Johnson, S. N., Douglas, A. E., Woodward, S. and Hartley, S. E. (2003). Microbial impacts on plant-herbivore interactions: the indirect effects of a birch pathogen on a birch aphid. Oecologia 134, 388–396. Jones, A. M. E., Bridges, M., Bones, A. M., Cole, R. and Rossiter, J. T. (2001). Purification and characterisation of a non-plant myrosinase from the cabbage aphid, Brevicoryne brassicae (L.). Insect Biochem. Mol. Biol. 31, 1–5. Jones, A. M. E., Winge, P., Bones, A. M., Cole, R. and Rossiter, J. T. (2002). Characterization and evolution of a myrosinase from the cabbage aphid Brevicoryne brassicae. Insect Biochem. Mol. Biol. 32, 275–284. Karley, A. J., Douglas, A. E. and Parker, W. E. (2002). Amino acid composition and. nutritional quality of potato leaf phloem sap for aphids. J. exp. Biol. 205, 3009–3018. Kehr, J., Hustiak, F., Walz, C., Wilmitzer, L. and Fisahu, J. (1998). Transgenic plants changed in carbon allocation pattern display a shift in diurnal growth pattern. Plant J. 16, 497–503. Kelly, P. J., Bones, A. M. and Rossiter, J. T. (1998). Subcellular immunolocalisation of the glucosinolate sinigrin in seedlings of Brassica junca. Planta 206, 370–377. Kempers, R. and van Bel, A. J. E. (1997). Symplasmic connections between sieve element and companion cell in the stem phloem of Vicia faba L have a molecular exclusion limit of at least 10 kDa. Planta 201, 195–201. Kennedy, J. S. and Mittler, T. E. (1953). A method for obtaining phloem sap via the mouth parts of aphids. Nature 171, 528. King, R. W. and Zeevart, J. A. D. (1974). Enhancement of phloem exudation from cut petioles by chelating agents. Plant Physiol. 53, 96–103.
THE NUTRITIONAL PHYSIOLOGY OF APHIDS
135
Klages, K., Donnison, H., Wu¨nsche, J. and Boldingh, H. (2001). Diurnal changes in non-structural carbohydrates in leaves, phloem exudates and fruit in ‘Braeburn’ apple. Austr. J. Pl. Physiol. 28, 131–139. Klingauf, F. A. (1987). Feeding, adaptation and excretion. In: Aphids Their Biology, Natural Enemies and Control (eds. Minks, A. K., and Harrewijn, P.) Vol. 1. pp. 225–253. Amsterdam: Elsevier. Kochian, L. V. (2000). Molecular physiology of mineral nutrient acquisition, transport and utilization. In: Biochemistry and Molecular Biology of Plants (eds. Buchanan, B., Gruissem, W., and Jones, R.), pp. 1204–1249. Rockville, MD, USA: American Society of Plant Physiologists. Ko¨ckenberger, W., Pope, J. M., Xia, Y., Jeffrey, K. R., Komor, E. and Callaghan, P. T. (1997). A non-invasive measurement of phloem and xylem water flow in castor bean seedlings by nuclear magnetic resonance microimaging. Planta 201, 53–63. Korslova, O. A., Davies, A., Deeken, R., Thorpe, M. R., Tomos, A. D. and Hedrich, R. (2000). Identification of a new glucosinolate-rich cell type in Arabidopsis flower stalk. Plant Physiol. 124, 599–608. Kunkel, H. and Hertel, R. (1975). Erstellung einer Stoffwechselbilanz fu¨r beide virginoparen Morphen von Myzus persicae (Sulz.) – Larven im dritten Stadium. Oecologia 18, 23–35. Lai, C-Y., Baumann, L. and Baumann, P. (1994). Amplification of trpEG: adaptation of Buchnera aphidicola to an endosymbiotic association with aphids. Proc. Natl. Acad. Sci. USA 91, 3819–3823. Lohaus, G., Hussmann, M., Pennewiss, K., Schneider, H., Zhu, J-J. and Sattelmacher, B. (2000). Solute balance of a maize (Zea mays L.) source leaf as affected by salt treatment with special emphasis on phloem retranslocation and ion leaching. J. Exp. Bot. 51, 1721–1732. Lucas, W. J. and Madore, M. A. (1988). Recent advances in sugar transport. In: The Biochemistry of Plants (eds. Preiss, J.) Vol. 14. pp. 35–84. Florida, Orlando: Academic Press. MacGibbon, D. B. and Allison, R. M. (1968). A glucosinolase system in the cabbage aphid, Brevicoryne brassicae. New Zealand J. Sci. 11, 440. McCauley, M. M. and Evert, R. F. (1989). Minor veins of the potato (Solanum tuberosum) leaf: ultrastructure and plasmodesmatal frequency. Bot. Gaz. 150, 351–368. Mady, E., Nowack, L. M. and Thompson, J. E. (2002). Isolation and characterization of lipid in phloem sap of canola. Planta 214, 625–634. Marschner, H., Kirkby, E. A. and Cakmak, I. (1996). Effect of mineral nutritional status on shoot-root partitioning of photoassimilates and cycling of mineral nutrients. J. Exp. Bot. 47, 1255–1263. Martin, B. J. L., Collar, W. F., Tjallingii, W. F. and Fereres, A. (1997). Intracellular ingestion and salivation by aphids may cause the acquisition and inoculation of non-persistently transmitted plant viruses. J. Gen. Virol. 78, 2701–2705. Matsiliza, B. and Botha, C. E. J. (2002). Aphid (Sitobion yakini) investigation suggests thin-walled sieve tubes in barley (Hordeum vulgare) to be more functional than thick-walled sieve tubes. Physiol. Planta. 115, 137–143. Merritt, S. Z. (1996). Within-plant variation in concentrations of amino acids, sugar, and sinigrin in phloem sap of black mustard, Brassica nigra (L.) Koch. J. Chem. Ecol. 22, 1133–1145. Miles, P. W. (1999). Aphid saliva. Biol. Rev. 74, 41–85. Mittler, T. E. (1958). Studies on the feeding and nutrition of Tuberolachnus salignus (Gmelin) (Homoptera Aphididae). II. The nitrogen and sugar composition of ingested phloem sap and excreted honeydew. J. Exp. Biol. 35, 74–84.
136
A. E. DOUGLAS
Mittler, T. E. (1971). Dietary amino acid requirement of the aphid Myzus persicae affected by antibiotic uptake. J. Nutr. 101, 1023–1028. Mittler, T. E. and Meikle, T. (1991). Effects of dietary sucrose concentration on aphid honeydew carbohydrate levels and rates of excretion. Entomol. Exp. Appl. 59, 1–7. Moericke, V. and Mittler, T. E. (1966). Oesophageal and stomach inclusions of aphids feeding on various Cruciferae. Entomol. Exp. Appl. 9, 287–297. Montalk, G. P., Remand-Simeon, M., Willemot, R-M., Sarcabal, P., Planchot, V. and Monsan, P. (2000). Amylosucrase from Neisseria polysacharea: novel catalytic properties. FEBS Lett. 471, 219–223. Montllor, C. B. (1991). The influence of plant chemistry on aphid feeding behaviour. In: Insect-Plant Interactions (eds. Bernays, E. A.) Vol. III. pp. 125–173. Boca Raton, Fl.: CRC Press. Montllor, C. B., Maxmen, A. and Purcell, A. H. (2002). Facultative bacterial endosymbionts benefit pea aphid Acyrthosiphon pisum under heat stress. Ecol. Entomol. 27, 189–195. Moran, N. A., Munson, M. A., Baumann, P. and Ishikawa, H. (1993). A molecular clock in endosymbiotic bacteria is calibrated using the insect hosts. Proc. Roy. Soc. London B 253, 167–171. Moran, P. J. and Thompson, G. A. (2001). Molecular responses to aphid feeding in Arabidopsis in relation to plant defense pathways. Plant Physiol. 125, 1074–1085. Munson, M. A., Baumann, P. and Kinsey, M. G. (1991). Buchnera gen. nov. and Buchnera aphidicola sp. nov., a taxon consisting of the mycetocyte-associated, primary endosymbionts of aphids. Int. J. Syst. Bac. 41, 566–568. Noiraud, N., Maurousset, L. and Lemoine, R. (2001). Transport of polyols in higher plants. Pl. Physiol. Biochem. 39, 717–728. Olaifa, J. I., Matsumura, F., Zeevaart, J. A. D., Mullin, C. A. and Charalambous, P. (1991). Lethal amounts of ricinine in green peach aphids (Myzus persicae) (Sulzer) fed on castor bean plants. Plant Sci. 73, 253–256. Passamani, M. and Rylands, A. B. (2000). Feeding behaviour of Geoffrey’s marmoset (Callithrix geoffroyi) in an Atlantic forest fragment of south-eastern Brazil. Primates 41, 27–38. Peel, A. J. (1975). Investigations with aphid stylets into the physiology of the sieve tube. In: Encyclopedia of Plant Physiology, New Series (eds. Zimmermann, M. H., and Milburn, J. A.) Vol. 1. pp. 171–195. Berlin: Springer Verlag. Percy, K. E., Awmack, C. S., Lindroth, R. L., Kubiske, M. E., Kopper, B. J., Isebrands, J. G., Pregitzer, K. S., Hendrey, G. R., Dickson, R. E., Zak, D. R., Oksanen, E., Sober, J., Harrington, R., Karnosky, D. F. (2002). Altered performance of forest pests under atmospheres enriched by CO2 and O3. Nature 420, 403–407. Pereira, S., Carvalho, H., Sunkel, C. and Salema, R. (1992). Immunolocalization of glutamine synthetase in mesophyll and phloem of leaves of Solanum tuberosum L. Protoplasma 167, 66–73. Pollard, D. G. (1973). Plant penetration by feeding aphids (Hempitera, Aphidoidea): a review. Bull. Ent. Res. 62, 631–714. Ponder, K. L., Pritchard, J., Harrington, R. and Bale, J. S. (2000). Difficulties in location and acceptance of phloem sap combined with reduced concentration of phloem amino acids explain lowered performance of the aphid Rhopalosiphum padi on nitrogen-deficient barley (Hordeum vulgare) seedlings. Entomol. Exp. Appl. 97, 203–210. Ponsen, M. B. (1979). The digestive system of Subsaltusaphis ornata (Homoptera: Aphididae). Mededelinger Landbouwhogeschol. Wageningen 79, 1–30. Ponsen, M. B. (1981). The digestive system of Eulachnus brevipilosus (Homoptera: Aphididae). Mededelinger Landbouwhogeschol. Wageningen 81, 1–14.
THE NUTRITIONAL PHYSIOLOGY OF APHIDS
137
Ponsen, M. B. (1987). Alimentary tract. In: Aphids: Their Biology, Natural Enemies and Control (eds. Minks, A. K., and Harrewijn, P.), pp. 79–97. Amsterdam: Elsevier. Prado, E. and Tjallingii, W. F. (1994). Aphid activities during sieve element punctures. Entomol. Exp. Appl. 72, 157–165. Prosser, W. A., Simpson, S. G. and Douglas, A. E. (1992). How an aphid (Acyrthosiphon pisum) symbiosis responds to variation in dietary nitrogen. J. Insect Physiol. 38, 301–307. Radford, J. E. and Overall, R. L. (2001). Effects of tissue-preparation-induced callose synthesis on estimates of plasmodesmata exclusion limits. Protoplasma 216, 47–55. Rahbe´, Y., Delobel, B., Febvay, G. and Chantegrel, B. (1994). Aphid specific triglycerides in symbiotic and aposymbiotic Acyrthosiphon pisum. Insect Biochem. Mol. Biol. 24, 95–101. Rahbe´, Y., Sauvion, N., Febvay, G., Peumans, W. J. and Gatehouse, A. M. R. (1995). Toxicity of lectins and processing of ingested proteins in the pea aphid Acyrthosiphon pisum. Entomol. Exp. Appl. 76, 143–155. Rahbe´, Y., Digilio, M. C., Febvay, G., Guillaud, J., Fanti, P. and Pennacchio, F. (2002). Metabolic and symbiotic interactions in amino acid pools of the pea aphid, Acyrthosiphon pisum, parasitized by the braconid Aphidius ervi. J. Insect Physiol. 48, 507–516. Raubenheimer, D. and Simpson, S. J. (1994). The analysis of nutrient budgets. Functional Ecology 8, 83–91. Raven, J. A. (1983). Phytophages of xylem and phloem – a comparison of animal and plant sap feeders. Adv. Ecol. Res. 13, 135–234. Rennenberg, H. (1982). Glutathione metabolism and possible biological roles in higher plants. Phytochem. 21, 2771–2781. Rhodes, J. D., Croghan, P. C. and Dixon, A. F. G. (1996). Uptake, excretion and respiration of sucrose and amino acids by the pea aphid Acyrthosiphon pisum. J. exp. Biol. 199, 1269–1276. Rhodes, J. D., Croghan, P. C. and Dixon, A. F. G. (1997). Dietary sucrose and oligosaccharide synthesis in relation to osmoregulation in the pea aphid, Acyrthosiphon pisum. Physiol. Entomol. 22, 373–379. Rispe, C. and Moran, N. A. (2000). Accumulation of deleterious mutations in endosymbionts: Muller’s ratchet with two levels of selection. Am. Nat. 156, 425–441. Rothschild, M. J., von Euw, J. and Reichstein, T. (1970). Cardiac glycosides in the oleander aphid Aphis nerii. J. Insect Physiol. 16, 1141–1145. Salvucci, M. E. and Crafts-Brandner, S. J. (2000). Effects of temperature and dietary sucrose concentration on respiration in the silverleaf whitefly, Bemisia argentifolii. J. Insect Physiol. 46, 1461–1467. Salvucci, M. E., Rosell, R. C. and Brown, J. K. (1998). Uptake and metabolism of leaf proteins by the silverleaf whitefly. Arch. Insect Biochem. Physiol. 39, 155–165. Sandstro¨m, J. (2000). Nutritional quality of phloem sap in relation to host plantalternation in the bird cherry-oat aphid. Chemoecol. 10, 17–24. Sandstro¨m, J., Telang, A. and Moran, N. A. (2000). Nutritional enhancement of host plants by aphids – a comparison of three aphid species on grasses. J. Insect Physiol. 46, 33–40. Sandstro¨m, J. P., Russell, J. A., White, J. P. and Moran, N. A. (2001). Independent origins and horizontal transfer of bacterial symbionts of aphids. Mol. Ecol. 10, 217–228. Sasaki, T., Aoki, T., Hayashi, H. and Ishikawa, H. (1990). Amino acid composition of the honeydew of symbiotic and aposymbiotic pea aphids Acyrthosiphon pisum. J. Insect Physiol. 36, 35–40.
138
A. E. DOUGLAS
Sasaki, T., Kawamura, M. and Ishikawa, H. (1996). Nitrogen recycling in the brown planthopper, Nilaparvata lugens: involvement of yeast-like endosymbionts in uric acid metabolism. J. Insect Physiol. 42, 125–129. Schuler, M. A. (1996). The role of cytochrome P450 monooxygenases in plant-insect interactions. Plant Physiol. 112, 1411–1419. Shigenobu, S., Watanabe, H., Hattori, M., Sakaki, Y. and Ishikawa, H. (2000). Genome sequence of the endocellular bacterial symbiont of aphids Buchnera sp. APS. Nature 407, 81–86. Silva, C. P., Ribiero, A. F., Gulbenkian, S. and Terra, W. R. (1995). Organisation, origin and function of the outer microvillar (perimicrovillar) membranes of Dysdercus peruvianus (Hemiptera) midgut cells. J. Insect Physiol. 41, 1093–1103. Silva, F. J., Latorre, A. and Moya, A. (2001). Genome size reduction through multiple events of gene disintegration in Buchnera APS. Trends Gen. 17, 615–618. Simpson, S. J. (1995). Regulation of a meal: chewing insects. In: Regulatory Mechanisms in Insect Feeding (eds. Chapman, R. F., and de Boer, G.), pp. 137–156. New York: Chapman and Hall. Simpson, S. J. and Raubenheimer, D. (1993). A multi-level analysis of feeding behaviour: the geometry of nutritional decisions. Phil. Trans. Roy. Soc. Lond. B 342, 381–402. Singer, M. S., Bernays, E. A. and Carrie`re, Y. (2002). The interplay between nutrient balancing and toxin dilution in foraging by a generalist and specialist insect herbivore. Anim. Behav. 64, 629–643. Spiller, N. J., Kimmins, F. M. and Llewellyn, M. (1985). Fine structure of aphid stylet pathways and its use in host plant resistance studies. Entomol. Exp. Appl. 38, 293–295. Srivastava, P. N. (1987). Nutritional physiology. In: Aphids Their Biology, Natural Enemies and Control (eds. Minks, A. K., and Harrewijn, P.) Vol. 1. pp. 99–122. Amsterdam: Elsevier. Srivastava, P. N. and Auclair, J. L. (1962). Characteristics of invertase from the alimentary canal of the pea aphid, Acyrthosiphon pisum (Harr.) (Homoptera: Aphididae). J. Insect Physiol. 9, 469–474. Srivastava, P. N. and Auclair, J. L. (1963). Characteristics and nature of proteases from the alimentary canal of the pea aphid, Acyrthosiphon pisum (Harr.) (Homoptera: Aphididae). J. Insect Physiol. 9, 469–474. Stadler, B. (1995). Adaptive allocation of resources and life history trade offs in aphids relative to plant quality. Oecologia 102, 246–254. Svoboda, J. A. and Feldlaufer, M. F. (1991). Neutral sterol metabolism in insects. Lipids 26, 614–618. Tamas, I., Klasson, L., Canback, B., Naslund, A. K., Eriksson, A. S., Wernegreen, J. J., Sandstrom, J. P., Moran, N. A. and Andersson, S. G. E. (2002). 50 million years of genomic stasis in endosymbiotic bacteria. Science 296, 2376–2379. Telang, A., Sandstro¨m, J., Dyreson, E. and Moran, N. A. (1999). Feeding damage by Diuraphis noxia results in nutritionally enhanced phloem diet. Entomol. Exp. Appl. 91, 403–412. Terra, W. R. (1990). Evolution of digestive systems of insects. Annu. Rev. Entomol. 35, 181–200. Terra, W. R., Ferreira., C. and Baker, J. E. (1996). Compartmentalisation of digestion. In: Biology of the Insect Midgut (eds. Lehane, M. J., and Billingsley, P. F.), pp. 206–235. London: Chapman and Hall. Thompson, G. A. and Schulz, A. (1999). Macromolecular trafficking in the phloem. Trends Pl. Sci. 4, 354–360. Tjallingii, W. F. (1995). Regulation of phloem sap feeding by aphids. In: Regulatory Mechanisms in Insect Feeding (eds. Chapman, R. F., and de Boer, G.), pp. 190–209. New York: Chapman and Hall.
THE NUTRITIONAL PHYSIOLOGY OF APHIDS
139
Tjallingii, W. F. and Esch, T. H. (1993). Fine structure of aphid stylet routes in plant tissues in correlation with EPG. Physiol. Entomol. 18, 317–328. Tomos, A. D. and Sharrock, R. A. (2001). Cell sampling and analysis (SiCSA): metabolites measured at single cell resolution. J. Exp. Bot. 52, 623–630. Turgeon, R. (1991). In ‘‘Recent Advances in Phloem Transport and Assimilate Compartmentation’’ (eds J.L. Bonnemain, S. Delrot, W.J. Lucas and J. Dainty), Symplastic phloem loading and sink-source transition in leaves: a model. pp. 98136. Ouest Editions, Nantes, France. Turgeon, R. and Medville, R. (1998). The absence of phloem loading in willow leaves. Proc. Natl Acad. Sci. 95, 12055–12060. Turgeon, R., Medville, R. and Nixon, K. C. (2001). The evolution of minor vein phloem and phloem loading. Am. J. Bot. 88, 1331–1339. Van Bel, A. J. E. and Gamalei, Y. V. (1992). Ecophysiology of phloem loading in source leaves. Plant Cell Env. 15, 265–270. Van Bel, A. J. E. and Knoblauch, M. (2000). Sieve element and companion cell: the story of the comatose patient and the hyperactive nurse. Aust. J. Plant Physiol. 27, 477–487. Van Bel, A. J. E., Ehlers, K and Knoblauch, M. (2002). Sieve elements caught in the act. Trends Plant Sci. 7, 126–132. Van Ham, R. C. H. J., Martinez-Torres, D., Moya, A. and Latorre, A. (1999). Plasmidencoded anthranilate synthase (TrpEG) in Buchnera aphidicola from aphids of the family Pemphigidae. Appl. Env. Microbiol. 65, 117–125. Van Helden, M. and Tjallingii, W. F. (1993). Tissue localisation of lettuce resistance to the aphid Nasonovia ribisnigri using electrical penetration graphs. Entomol. Exp. Appl. 68, 269–278. Volk, G. M. and Franceschi, V. R. (2000). Localisation of a calcium channel-like protein in the sieve element plasma membrane. Aust. J. Plant Physiol. 27, 779–786. Von Dohlen, C. D. and Moran, N. A. (1995). Molecular phylogeny of the Homoptera: a paraphyletic taxon. J. Mol. Evol. 41, 211–223. Vrieling, K., Smit, W. and van der Meijden, E. (1991). Tritrophic interactions between aphids (Aphis jacobaea Schrank), ant species, Tyria jacobaeae L., and Senecio jacobaea L. lead to maintenance of genetic vairiation in pyrrolizidine alkaloid concentration. Oecologia 86, 177–182. Waller, G. R. and Skursky, L. (1972). Translocation and metabolism of ricinine in the castor bean plant, Ricinus communis L. Plant Physiol. 50, 622–626. Walling, L. L. (2000). The myriad plant responses to herbivores. J. Pl. Growth Regul. 19, 195–216. Walters, F. S. and Mullin, C. A. (1988). Sucrose-dependent increase in oligosaccharide production and associated glycosidase activities in the potato aphid Macrosiphum euphorbiae (Thomas). Arch. Insect Biochem. Physiol. 9, 35–46. Walters, F. S., Mullin, C. A. and Gildow, F. E. (1994). Biosynthesis of sorbic acid in aphids – an investigation into symbiont involvement and potential relationship with aphid pigments. Arch. Insect Biochem. 26, 49–67. Ward, J. M., Ku¨hn, C., Tegeder, M. and Frommer, W. B. (1998). Sucrose transport in higher plants. Int. Rev. Cytol. 178, 41–71. Weibull, J., Ronquist, F. and Brishammar, S. (1990). Free amino acid composition of leaf exudates and phloem sap. A comparative study in oats and barley. Plant Physiol. 92, 222–226. Wellings, P. W. and Dixon, A. F. G. (1983). Physiological constaints on the reproductive activity of the sycamore aphid: the effect of developmental experience. Entomol. Exp. Appl. 34, 227–232. Wernegreen, J. J., Ochman, H., Jones, I. B. and Moran, N. A. (2000). Decoupling of genome size and sequence divergence in a symbiotic bacterium. J. Bac. 182, 3867–3869.
140
A. E. DOUGLAS
Wernegreen, J. J., Richardson, A. O. and Moran, N. A. (2001). Parallel acceleration of evolutionary rates in symbiont genes underlying host nutrition. Mol. Phylogenet. Evol. 19, 479–485. Whitehead, L. F. (1993). The Symbiotic Bacteria of the Pea Aphid, Acyrthosiphon pisum: Ph.D. Thesis, University of Oxford, UK. Whitehead, L. F., Wilkinson, T. L. and Douglas, A. E. (1992). Nitrogen recycling in the pea aphid (Acyrthosiphon pisum) symbiosis. Proc. Roy. Soc. Lond. B 250, 115–117. Whitehead, L. F. and Douglas, A. E. (1993a). Populations of symbiotic bacteria in the parthenogenetic pea aphid (Acyrthosiphon pisum) symbiosis. Proc. Roy. Soc. London B 254, 29–32. Whitehead, L. F. and Douglas, A. E. (1993b). A metabolic study of Buchnera, the intracellular bacterial symbionts of the pea aphid, Acyrthosiphon pisum. J. Gen. Microbiol. 139, 821–826. Wilkinson, T. L. and Douglas, A. E. (1995). Why aphids lacking symbiotic bacteria have elevated levels of the amino acid glutamine. J. Insect Physiol. 41, 921–927. Wilkinson, T. L. and Douglas, A. E. (2003). Phloem amino acids and the host plant range of the polyphagous aphid, Aphis fabae. Entomol. Exp. Appl. 106, 1–11. Wilkinson, T. L. and Ishikawa, H. (1999). The assimilation and allocation of nutrients by symbiotic and aposymbiotic pea aphids, Acyrthosiphon pisum. Entomol. Exp. Appl. 91, 195–201. Wilkinson, T. L. and Ishikawa, H. (2000). Injection of essential amino acids substitutes for bacterial supply in aposymbiotic pea aphids (Acyrthosiphon pisum). Entomol. Exp. Appl. 94, 85–91. Wilkinson, T. L., Ashford, D. A., Pritchard, J. and Douglas, A. E. (1997). Honeydew sugars and osmoregulation in the pea aphid Acyrthosiphon pisum. J. Exp. Biol. 200, 2137–2143. Wilkinson, T. L., Adams, D., Minto, L. B. and Douglas, A. E. (2001). The impact of host plant on the abundance and function of symbiotic bacteria in an aphid. J. Exp. Biol. 204, 3027–3038. Williams, C. T. (1995). Effect of plant age and condition on the population dynamics of Myzus persicae Sulz. on sugar beet in field plots. Bull. Entomol. Res. 85, 557–567. Williams, I. S., Dewar, A. M. and Dixon, A. F. G. (1997). The effect of host plantinduced stomach precipitate on the ability of Myzus persicae (Hemiptera: Aphididae) to transmit sugarbeet yellowing viruses. Bull Entomol. Res. 87, 643–647. Wink, M. and Witte, L. (1991). Storage of quinolizidine alkaloids in Macrosiphum albifrons and Aphis genistae (Hompotera, Aphididae). Entomol. Gen. 15, 237–254. Winter, H., Lohaus, G. and Heldt, H. W. (1992). Phloem transport of amino acids in relation to their cytosolic levels in barley leaves. Plant Physiol. 99, 996–1004. Wolfe, G. R., Hendrix, D. L. and Salvucci, M. E. (1998). A thermoprotective role for sorbitl in the silverleaf whitefly, Bemisia argentifolii. J. Insect Physiol. 44, 597–603. Wolfe, G. R., Smith, C. A., Hendrix, D. L. and Salvucci, M. E. (1999). Molecular basis for thermoprotection in Bemisia: structural differences between whitefly ketose reductase and other medium chain dehydrogenases/reductases. Insect Biochem. Mol. Biol. 29, 113–120. Wolfersberger, M. G. (2000). Amino acid transport in insects. Annu. Rev. Entomol. 45, 111–120. Wright, J. P., Fisher, D. B. and Mittler, T. E. (1985). Measurement of aphid feeding rates of artificial diets using 3H-inulin. Entomol. Exp. Appl. 37, 9–11. Ziegler, H. (1975). Nature of transported substances. In: Encyclopedia of Plant Physiology, New Series (eds. Zimmermann, M. H., and Milburn, J. A.) Vol. 1. pp. 59–100. Berlin: Springer Verlag.
The Neurobiology of Taste in Insects Stephen M. Rogersa and Philip L. Newlandb a Department of Zoology, University of Cambridge, Downing St, Cambridge CB2 3EJ, UK; b School of Biological Sciences, University of Southampton, Bassett Crescent East, Southampton SO16 7PX, UK
1 Introduction 142 1.1 Components of insect chemosensory systems 144 1.2 Sensilla locations and behavioural hierarchies 144 2 Chemosensory coding 147 2.1 Olfactory coding 148 2.2 Potential difficulties with gathering and interpreting contactchemosensory neurone responses 149 2.3 Different hypotheses about contact-chemosensory coding 150 2.4 Variation in chemosensory response 156 3 Central projections of sensory neurones 159 3.1 Central organization of sensory neurones from tactile hairs 160 3.2 Central organization of sensory neurones from contact-chemosensory sensilla 162 3.3 Modality-specific segregation 164 3.4 Somatotopic mapping of sensory neurones innervating contactchemosensory sensilla 167 3.5 Chemosensory mapping 168 4 Local circuits and their role in processing gustatory signals 170 4.1 Processing of sensory signals 172 4.2 The motor output of local circuits 179 5 Chemosensory coding in the metathoracic ganglion of the locust 180 5.1 Behavioural responses to chemosensory stimulation 180 5.2 Responses of spiking local interneurones to different chemical solutions 182 5.3 Responses of leg motor neurones 185 5.4 Significance and general applicability of the locust model 188 6 Concluding remarks 192 Acknowledgements 193 References 194
ADVANCES IN INSECT PHYSIOLOGY VOL. 31 ISBN 0-12-024231-1 DOI: 10.1016/S0065-2806(03)31003-3
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Abstract Taste or contact-chemoreception is a fundamentally important sense to most insects. Despite its importance there remains a longstanding controversy about how different contact-chemosensory qualities are coded in the central nervous system and several different models have been proposed. Sensory projections from contact-chemosensory sensilla are primarily to the local segmental ganglion or neuromeres. There appears to be a distinction between Orthopteroid insects in which chemosensory and mechanosensory neurones project to the same, somatotopically defined, regions of neuropile, and the Diptera where there is evidence for a modality-specific spatial separation of neurones. Recent advances have been made in understanding central contactchemosensory processing using the hind leg-metathoracic ganglion of the locust as a model system. This system offers the advantage of defined local behavioural responses controlled by well-understood local neuronal networks. Chemosensory and mechanosensory neurones make monosynaptic connections onto the same spiking local interneurones. All chemicals can elicit withdrawal responses of a leg but the concentration at which different chemicals become effective stimuli varies over several orders of magnitude. The relative size of response of local interneurones to these same chemicals, as well as their outputs onto leg motor neurones, is closely correlated with the probability of eliciting a behavioural withdrawal response. We suggest that contact-chemosensory processing by local circuits in the thoracic ganglia directly assesses a chemosensory quality, that of aversiveness, which is dependent on both chemical identity and concentration and that other chemical qualities such as palatability may be encoded in a similar manner elsewhere in the central nervous system.
1
Introduction
Feeding is one of the most complex functions an animal must perform, involving the coordinated activity of many neuronal networks, integration from several sensory modalities and the organization of complex patterns of motor activity in even the simplest insect (Kupfermann, 1994). The sense of taste or contact-chemoreception has a fundamental role in the process of feeding in many insects. In this review we intend to highlight some of the recent advances in our understanding of the neurobiology of taste in insects in both the peripheral and central nervous systems, ranging from chemosensory coding during initial food selection and how chemosensory variability may be used to regulate dietary intake, through to the processing of contact-chemosensory information in the central nervous system and the organization of appropriate
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behavioural responses. We will also outline some of the remaining problems that have yet to be fully resolved. Since insect taste receptors are readily accessible, and their sensory activity readily monitored, studies of the chemosensory basis of food detection and selection have dominated investigations into the neurobiology of feeding in insects. While it has been acknowledged that the visual system may have an important role in host plant selection (Bell, 1990; Wacht et al., 1996), this role has yet to be systematically investigated at a neurobiological level. This contrasts with what we know of the visual targeting of prey by carnivorous insects such as mantids (e.g. Kral, 1998; Gonka et al., 1999; Kral and Devetak, 1999). Olfaction in insects has been extensively analysed and it is not our intention here to cover olfactory processing in detail since several recent reviews cover the subject more judiciously than we could hope to achieve (for example, Hildebrand and Shepherd, 1997; Hansson and Anton, 2000; Laurent et al., 2001; Christensen and Hildebrand, 2002). We will refer to olfactory processing only in so far as it contrasts with contact chemosensory, or gustatory, coding and/or is explicitly involved in food selection. Contact-chemoreception has a central role in the life of insects. Tasks such as finding and assessing the qualities of potential foods, avoiding noxious or otherwise harmful substances in the environment, finding mates and suitable oviposition sites would be impossible for most insects without welldeveloped contact chemosensory systems. These same chemosensory systems are also involved intimately in the regulation of feeding behaviour, not only allowing an insect to reject unacceptable foods but also to dynamically regulate the intake of potentially acceptable foods in the light of its current nutritional requirements, and also in helping to determine when to feed and for how long. The rules governing food selection and dietary regulation in insects have been extensively investigated at the behavioural level in several species of insects and contact-chemosensory information is thought to make an important contribution to the processes underlying these behaviours (Dethier, 1976; Simmonds et al., 1992; Simpson, 1994; Raubenheimer and Simpson, 1993; Simpson and Raubenheimer, 1993a,b, 1996; Amakawa, 2001). The peripheral sensory physiology of the chemosensory systems has been extensively analysed in many insect species. Whilst considerable progress has been made in understanding the organization and some of the neural interactions underlying olfactory processing (although it is far from being completely understood), next to nothing is known of the central integration of gustatory information, which potentially has the greater and more immediate role in dietary selection and the control of feeding. Part of the argument we wish to develop is to suggest a functional distinction between olfaction and gustation that depends upon the features of the chemosensory environment that each sense abstracts from a stimulus. Thus although ‘gustatory’ receptors
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have in some instances been shown to respond to certain odours, i.e. a chemosensory stimulus in a vapour phase (Sta¨dler and Hanson, 1975; Newland, 1998), it is likely that the central nervous system processes these stimuli in a very different way from the same odour sampled by classical ‘olfactory’ receptors and centrally processed by the antennal lobes of the brain. We will outline evidence from our recent experiments analysing taste processing in locusts that contact-chemoreception and mechanoreception are closely related to each other and that these two modalities are processed, at least in part, by the same pathways in the central nervous system. The common occurrence of chemosensory and mechanosensory neurones within the same sense organ therefore reflects the co-processing of these two modalities. 1.1
COMPONENTS OF INSECT CHEMOSENSORY SYSTEMS
Insect chemoreceptors are primary afferent neurones that are contained within cuticular structures called sensilla and these, along with supporting cells (Fig. 1), constitute the basic organ of chemoreception in arthropods (Lewis, 1970; Zacharuk, 1980, 1985). Externally, chemosensory sensilla are extremely diverse in structure across insect groups, and sometimes even between different locations on the same insect, but most frequently take the form of small hollow hairs or pegs bearing one or more pores through to their interior. Chemosensation may be mediated by tens to thousands of these structures depending on the species of insect (Schoonhoven, 1973; Chapman and Thomas, 1978; Chapman, 1982) and anything from two to fifty chemosensory neurones may be present in each sensillum. The division between uniporous sensilla, containing chemosensory dendrites that are simple unbranched rods, and multiporous sensilla, whose chemosensory dendrites are highly branched, matches, albeit imperfectly in some cases, the sub-division of chemoreception into the senses of gustation (or contactchemoreception) and olfaction (Chapman, 1982, 1995; Zacharuk, 1985; Blaney and Simmonds, 1990a). Uniporous sensilla are frequently attached to the cuticle by a flexible socket and typically also contain a single mechanosensory neurone as well as chemosensory neurones. Therefore, these sensilla mediate both taste and touch. General reviews of the ultrastructure of insect chemosensory sensilla can be found in Altner (1977) and Zacharuk (1985). 1.2
SENSILLA LOCATIONS AND BEHAVIOURAL HIERARCHIES
Contact-chemosensory sensilla may be found distributed over the whole body surface in many insects, but usually they form local aggregations that define sensory fields located in key positions on the body consisting of few to hundreds of sensilla. These sensory fields are most commonly located on the tarsi, antennae, maxillae, labium, labrum and cibarial cavity. The total number of contact-chemosensory sensilla present on an insect varies enormously,
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FIG. 1 Cross-section through a typical contact-chemosensory sensillum. All sensory neurones and supporting cells originate from a single epithelial cell. The outer support cells (trichogen cells) secrete the shaft of the sensillum and hollow out to form the sensillum sinus. The wall of the sensillum sinus is highly convoluted and secretes the aqueous sensillum lymph as well as chemical binding proteins into the lumen. The inner support cells (tormagen cells), wrap around the somata of the sensory neurones and extend upwards into the shaft of the sensillum to form the dendrite sinus. Contactchemosensory sensilla typically have a single apical pore and contain several chemosensory neurones whose dendrites project to near the pore opening. A single mechanosensory neurone is a common component of these sensilla; its dendrite inserts into the tubular body attached to the flexible shaft of the sensillum.
ranging from tens to thousands, depending on the species, dietary preference, size, age and sex of the insect (Chapman, 1982). Caterpillars (Lepidoptera) and true bugs (Hemiptera) characteristically possess few contact-chemosensory sensilla, which are on key locations on the tarsi, antennae and mouthparts and the numbers of which do not increase with successive larval moults. The orthopteroid insects and beetles (Coleoptera) by contrast typically have large and variable numbers of sensilla in each of their sensory fields. In the orthopteroid orders the number of sensilla increases with each instar, and the total number is correlated with the overall size of the insect species. There is also a correlation between the number of sensilla and the dietary habit. Scavenging insects have the most, followed by polyphagous herbivores with oligophages and monophages having
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progressively fewer sensilla (Chapman and Fraser, 1989). Even monophagous orthopteroid insects, however, still possess several hundred chemosensory sensilla in total. The Diptera generally fall somewhere between these two extremes (Chapman, 1982), possessing fairly large numbers of chemosensory sensilla on their legs, but there is a population of large chemosensory sensilla on the labellum that are sufficiently low in numbers that in many cases at least some of the individual sensilla can be identified from animal to animal (Stocker, 1994; Edgecomb and Murdock, 1992). The antennae are dominated by multiporous chemosensilla and these form the primary site of olfaction in insects. Uniporous sensilla are most common on the general body surface and mouthparts. There are several known examples of both sensilla types occurring together within the same population. Uniporous sensilla are frequently present on the antennae, but usually in smaller numbers (Chapman, 1982; Greenwood and Chapman, 1984) and multiporous sensilla may be found on the external mouthparts (Blaney, 1977; Devitt and Smith, 1982). These may constitute fairly extensive secondary olfactory fields and can be found in several types of insects, for example, the maxillary palps in Diptera (Naresh Singh and Nayak, 1985; Distler and Boeckh, 1997), and caterpillars (Devitt and Smith, 1982) and the CO2detecting labial-palp pit organ in adult Lepidoptera (Bogner et al., 1986; Lee and Altner, 1986). Regardless of the total number of sensilla, it is almost invariably the case that whereas olfactory receptors on the antennae (Homberg et al., 1989) and most likely on the mouthparts all project to well-defined olfactory neuropils in the deuterocerebrum of the brain (Kent and Hildebrand, 1987; Rogers, 1998; de Bruyne et al., 1999), the widely dispersed contact-chemosensory neurones project to their local ganglion and there is no single specific gustatory processing region within the CNS (see Section 3). There is a hierarchy of chemosensory fields with respect to the order in which they usually encounter chemicals. Clearly the senses of olfaction and vision that are able to sample the environment from a distance have a critical role in the initial attraction of an insect to potential foods (Visser, 1986). Olfaction may also have an important and sustained role in the more intimate assessment of food quality that occurs on alighting on a potential food source, but it is at this stage that two new senses, those of contact-chemoreception and mechanoreception come into play. During typical food location behaviour, tarsal chemoreception is usually the first of a series of contact-chemosensory fields with which a potential food is assessed before ingestion occurs (Dethier, 1976; Simpson, 1992). Tarsal chemosensory cues may also be important in eliciting oviposition behaviour in insects searching for suitable host plants (Blaney and Simmonds, 1990b; Roessingh et al., 1991). The rapid withdrawal of the leg may follow the detection of aversive chemicals (Slifer, 1956; White and Chapman, 1990; Rogers and Newland, 2000). If the potential food is not rejected at this stage, and the tarsi detect suitable phagostimulatory cues, the
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head is usually lowered so that other contact-chemosensory fields located on the external mouthparts can sample it (Bernays and Simpson, 1982; Chapman, 1995; van Loon, 1996). The chemosensory sensilla in these fields frequently exhibit a greater sensitivity than those on the tarsi (Clements, 1992), and are commonly located on the tips of the maxillary and labial palps of orthopteroid insects (Blaney and Chapman, 1970; Mordue, 1979), the galea and maxillary palps of larval Lepidoptera (Schoonhoven and van Loon, 2002) and on the labellum of flies (Dethier, 1976). At this stage the antennae may be lowered and contact the surface. If the potential food is still not rejected the insect will usually proceed to biting the item. This is the first stage at which the internal contents of a food item, such as a leaf, will be accessible in high concentration to the chemosensory apparatus of the insect. Even intact leaves, however, normally leak their internal chemical contents and therefore the insect can gain some information about a leaf’s internal characteristics, albeit at a very low concentration, from sampling its surface prior to biting (Fiala et al., 1990; Derridj et al., 1996). The contact-chemosensory fields sampling bitten material are commonly located on the inner surface of the labrum and cibarial cavity (Chapman, 1982). If the item is still not rejected, it will be ingested and the insect will start feeding (Bernays and Simpson, 1982). This behavioural chain is not fixed since an item may be rejected at any stage, and the observation by White and Chapman (1990) of a locust holding its front tarsi clear of a nicotine hydrogen tartrate coated leaf whilst at the same time palpating and feeding on it, is evidence that rejection by one of the sensory fields need not always lead to the overall rejection of the food item. Nevertheless, if a potential food encountered by a naı¨ ve locust is rejected after biting, it is subsequently much more likely to be rejected at the earlier palpation stage, suggesting a degree of learned association between the experiences of the different sensory fields (Blaney and Winstanley, 1982; Blaney and Simmonds, 1985; Blaney et al., 1985). Hammer (1993) demonstrated a learned proboscis extension-reflex in bees, in which an association was formed between particular odours and the application of sugar solutions to the proboscis. This appears to be mediated by a single large neurone with branches in the brain and sub-oesophageal ganglion, though it is likely to be just one element in a more complex neuronal circuit. This demonstrates not only a close link between the olfactory and gustatory systems at both the behavioural and neural levels (Feeny et al., 1989; de Boer, 1993) but also the flexibility of the chain of chemosensory sampling behaviour.
2
Chemosensory coding
Our understanding of gustatory processing in insects has been governed almost entirely by the analysis of the activity patterns of sensory neurones to different
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sapid stimuli. Until recently there has been very little known about the destinations of contact-chemosensory neurones and the central processing of the gustatory information they carry. The central processing of contactchemoreception is probably one of the least investigated senses in insects, and stands in contrast with the wealth of work that has been done on the central processing of olfaction (Homberg et al., 1989; Masson and Mustaparta, 1990; Hildebrand, 1995, 1996). This situation has understandably led to two still unresolved debates: first, exactly what kind of information is carried by contactchemosensory neurones, whether chemical identity or a more abstract gustatory quality, and second how this information is actually represented by a population of chemosensory neurones. 2.1
OLFACTORY CODING
Before considering gustatory coding, it is worth briefly summarizing the state of knowledge about olfactory coding, to provide a point of reference against which different models of gustatory processing can be compared and contrasted. Whilst it seems likely that insects can recognize individual odours, possibly substances never previously encountered and of no immediate fitness consequence, and make learned associations between them and other stimuli (Tully, 1987; Brandes et al., 1988; Simpson and White, 1990; de Jong and Kaiser, 1991; de Jong and Pham-Delegue, 1991; Hammer, 1993; Smith and Getz, 1994), it is much less certain how gustatory information is represented in the CNS (Chapman, 1988, 1995). The ability of an insect to recognize individual odours is not perfect. Compounds chemically similar to the odorant originally used in learning procedures may be able to elicit the same conditioned response (Smith, 1991). In addition, population receptor responses may be much stronger to certain classes of odorant that are important to the insect, such as those associated with host plants or other foods (Nottingham et al., 1991; Wibe et al., 1996). The extreme of the trend towards olfactory foci for behaviourally important classes of chemicals is found in the segregated detecting and processing systems for pheromones. Other than pheromone receptors, most individual olfactory neurones will respond to a range of different odours. Each distinct odour will elicit a different pattern of activity in the chemoreceptor population, with some neurones responding and others not. Different odours evoke distinct, but overlapping activity-spectra and it is thought that individual odours are recognized through interpreting these patterns of activity in the chemoreceptor population as a whole, a process termed cross-fibre (or across-fibre, or vector) coding (Pfaffmann, 1941; Selzer, 1984; Boeckh and Ernst, 1987; Smith and Getz, 1994). However, where an odour consists of a blend of different molecules and two or more of these components excite the same sensory neurone, or sometimes even different neurones within the same sensillum, complex synergistic or inhibitory interactions may occur, so that the receptor
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response is not the simple sum of the responses to the individual components (Akers and Getz, 1993; Getz and Akers, 1994). Different odours on average activate approximately 10% of the total antennal olfactory receptor neurone array in locusts, irrespective of whether they are monomolecular or composed of a blend of different molecules (Laurent, 1996). This suggests that olfactory integration and discrimination is considerably more complex than a simple process of matching different molecular templates by the CNS (Getz and Akers, 1997). 2.2
POTENTIAL DIFFICULTIES WITH GATHERING AND INTERPRETING CONTACT-CHEMOSENSORY NEURONE RESPONSES
A wider diversity of models has been proposed to explain contactchemosensory coding than olfaction, a diversity helped because in most cases the only information available is the primary sensory response measured at the periphery. It is first worth considering the methods used to record contactchemosensory responses and some of the possible problems encountered when doing so. Chemosensory afferents are small and in their natural context almost completely inaccessible to intracellular recording techniques, with the consequence that many of the neuronal conductances that occur prior to the production of action potentials are still imperfectly understood. This understanding is essential to a deeper understanding of both transduction and of the modification of chemosensory responses, for example in the presence of mixtures (see Section 2.3). Nearly all recordings are extracellular, made either using the tip recording technique (Hodgson et al., 1955), or by sidewall or basal recording obtained by inserting sharp electrodes under the cuticle adjacent to or within the walls of a sensillum. A vast amount of information has been gathered from many types of sensilla on many species using these methods (e.g. see Schoonhoven and van Loon (2002), on caterpillars and Chapman (2003), for comprehensive reviews) but they have limitations. In tip recording all test chemicals must be in an aqueous electrolyte, usually dilute potassium or sodium chloride solution. In fluid-feeding insects these solutions may provide a reasonably close approximation of food substances but it is unclear how these solutions equate to the chemosensory environment found on the surfaces of solid matter, such as a leaf surface. It is likely that even internally leaf sap is not a homogeneous substrate but will contain reservoirs of high and low concentration. Moreover, it is difficult to test responses to hydrophobic substances, which again may be abundant in the waxes on a leaf surface and hence may form an important source of gustatory information (Sta¨dler, 1986; Woodhead and Chapman, 1986). Another considerable problem is how to adequately characterize a chemosensory neurone. At a practical level there is a limit to the number of chemical solutions that can be applied to a sensillum and our understanding
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of the key features of a ligand molecule that signal its acceptability to a particular receptor is still at an elementary stage (Araneda et al., 2000). A generalist chemosensory neurone is potentially able to respond to hundreds of compounds, and when variability in sensitivity is added to the problem, the range of concentrations at which these different compounds become effective stimuli of a chemosensory neurone is also potentially large. This often means that the experimenter’s judgement must be used as to what is likely to be an important stimulus and at what concentration it is likely to be effective. Therefore, despite the wealth of chemosensory response data in the literature, this is often fragmentary; different substances are tested at different concentrations on different sensilla on different species. Frequently, we simply do not know what the true capacity of a particular chemosensory neurone, or chemosensory sensillum, or group of sensilla is and how this compares between species. Needless to say this makes any appreciation of general coding principles of contact-chemosensation difficult. Schoonhoven and van Loon (2002) have recently produced a comprehensive review of known stimulants of various contact-chemosensory neurones in many species of Lepidoptera. When taken together some common themes emerge, but as the authors acknowledge in their title ‘each species its own key’ (Schoonhoven and van Loon 2002), what stands out is the diversity of chemosensory neurone responses between different species, a diversity perhaps exaggerated by a diversity of experimental techniques and stimuli.
2.3
DIFFERENT HYPOTHESES ABOUT CONTACT-CHEMOSENSORY CODING
Having established that there are problems in the collection and interpretation of data from contact-chemosensory sensilla, we will next consider existing hypotheses for how such inputs are processed. Suggestions have been made that range from across-fibre, fully discriminatory systems, akin to those used in olfaction (but perhaps simpler) (Blaney, 1975; Dethier and Crnjar, 1982; Van Loon, 1996; Glendinning et al., 2002); to labelled lines for specific phagostimulatory or inhibitory compounds (Du et al., 1995); to non-specific systems that assess the total phago-stimulatory and/or deterrent quality (Blaney and Winstanley, 1980; Blaney, 1981; Schoonhoven, 1987; Simpson and Raubenheimer, 1996), or combinations of the above. Early work on flies and caterpillars lead to the characterization of individual neurones within a sensillum as being tuned to respond to sugars, high concentrations of salts, or low concentrations of salts (the ‘water’ cell) (Ma, 1972; Schoonhoven, 1973; Dethier, 1976). Another neurone type found in caterpillars, characterized as the deterrent or ‘D’ neurone (Schoonhoven et al., 1992; Schoonhoven and van Loon, 2002), responds to secondary plant compounds and its activity is correlated with decreased feeding or aversion responses by the insect. A similar neurone has been described in acridids
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(Chapman et al., 1991). However, like olfactory neurones, individual gustatory sensory neurones are commonly responsive to a wide range of compounds, which may not be chemically similar. Locusts for example do not even appear to have distinct salt and sugar responsive neurones, as at least two neurones in each sensillum respond to both class of chemicals (Blaney, 1975). Given that, as discussed above, few chemosensory neurones have been fully chemically characterized, such labels may be at least incomplete descriptions or even misleading (Simmonds and Blaney, 1990). The recent identification of candidate gustatory receptor proteins (Clyne et al., 2000; Dunipace et al., 2001; Robertson, 2001) may eventually resolve some issues of the physiological range of individual chemosensory neurones. The isolation of these receptor proteins potentially allows for a more focused analysis of the critical features of their ligand molecules and resolve whether the wide-ranging sensitivity to chemically diverse stimuli reported for some insect gustatory neurones (Bernays et al., 2000) is itself the property of individual receptor proteins or arises from the presence of different receptor types within one neurone. Glendinning and co-workers (Glendinning et al., 1999, 2001, 2002) have, for instance, provided strong physiological evidence that individual deterrent sensory neurones in Manduca sexta possess two different receptor types that activate distinct transduction pathways. Contact-chemosensory neurones closely resemble those of olfactory receptors in their basic functioning and there has been a tacit assumption amongst many authors that individual chemicals can be fully discriminated between central neurones reading off the population response of chemosensory neurones using across-fibre coding (e.g. Blaney, 1975; Dethier and Crnjar, 1982; van Loon, 1996). In this model, contact-chemoreception is similar to olfaction, differing from it only in the range and type (e.g. non-volatile) of chemicals that can be detected. A potential problem arises with across-fibre coding when non-volatile stimuli are detected using chemosensory neurones contained within spatially separate sensilla. Not all the necessary neurones may be brought into contact onto an irregular surface and therefore the central nervous system may receive an erroneous population response. The common occurrence of a mechanosensory neurone in contact-chemosensory sensilla could provide the insect with the necessary information to prevent this happening. Another apparent similarity with the olfactory system is that different sensilla within a sensory field often contain neurones with different sensitivities to particular chemicals and/or respond to different chemicals (Blaney, 1975, 1981). Whilst these features are consistent with some form of across-fibre patterning, they could equally be interpreted as part of a non-discriminatory system in which inputs from sensory neurones, carrying either phago-stimulatory or deterrent information, are summed from several or many sensilla, some of which may contribute a different
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weighting towards the overall sensory input. Blaney (1981) performed an analysis of this sort on the responses of contact-chemosensory neurones in two species of locust to a number of different chemical stimuli. The total numbers of action potentials across all the chemosensory neurones within individual basiconic sensilla were merely weighted against the response of the same sensilla to electrolyte solution controls. This total ‘relative sensory input’ was correlated with the relative palatability of the stimuli as determined in behavioural assays, which measured the consumption of paper discs impregnated with the tested chemicals. In Schistocerca gregaria there was a strong and simple correlation between the unpalatability of a chemical and the total firing rate of chemosensory neurones across the population of sensilla, suggesting that Schistocerca will eat anything that does not elicit too strong a sensory response, a result which accords with our own analyses of the central processing of chemosensory stimuli (Rogers and Newland, 2002) that we describe in Section 5 below. In Locusta migratoria the relationship between palatability and overall sensory input was not as clear-cut. Indeed, Blaney (1981) suggested that overall, palatability was negatively correlated with total sensory input across a range of secondary plant compounds and positively correlated for sucrose, and that Locusta therefore makes more complex assessments of phagostimulatory and phagodeterrent sensory inputs than Schistocerca. Nevertheless, for both species both unpalatability and relative sensory input increased approximately linearly for any one chemical; Locusta differed from Schistocerca in that it lacked a simple all encompassing relationship between sensory input and unpalatabilty across all the chemicals tested, but within concentration series of any one chemical there was in most cases a simple positive correlation between sensory input and unpalatabilty. This may indeed point to a greater complexity of central processing, or it may more simply be explicable by differences in the synaptic weighting of different sensory neurones onto central chemosensory processing interneurones. Recently, Bernays and Chapman (2001) and Chapman (2003) have proposed another system of contact-chemosensory processing based on the direct assessment of chemical qualities rather than via chemical identity. They have based their model on an analysis of the chemosensory responses of two important sensilla on the galea of the caterpillar Grammia genura, correlated with behavioural responses to several chemical stimuli. They contend that each sensillum contains two general phagostimulatory and two general phagodeterrent neurones, activity in which are correlated with either food acceptance or rejection respectively. This system is non-discriminatory in that activity in each of the cells is generically concerned with either food acceptance or rejection irrespective of the chemical or chemicals stimulating the neurones. Their approach contrasts with earlier work in that it attempts to categorize chemosensory neurones by the behavioural effect they elicit rather than by the chemicals they are sensitive to.
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Simpson and Raubenheimer (1996) have developed a model of taste that is similarly not dependent upon an across-fibre-patterning/individual discrimination mode of central processing but nevertheless attempts to explain how the combined population response or gestalt of gustatory information reaching the central nervous system can be used by an insect to make a complex assessment of the quality of a potential food. In this model it is reasoned that the contactchemosensory system of an insect should have evolved to be tuned such that the foods giving the strongest phagostimulatory signal to the central nervous system should be those that most closely meet the insect’s nutritional requirements at that time. This is dependent not only on the chemicals present (or not present) in that food, but on their amounts, both in absolute quantities and relative to other chemicals or nutrients, and on the current nutritional state of the insect. This idea is allied to the concept of an intake target (Raubenheimer and Simpson, 1993), the quantity and blend of nutrients that an insect needs to perform optimally. This optimum may be expected to vary with developmental stage and previous dietary experience, and hence it reflects the future requirements of an insect to achieve a balanced and optimal diet. Therefore individual chemicals are not treated as sign stimuli, whose presence in a food means that it is always intrinsically appetising. Indeed high concentrations of single chemicals, even nutrients, can impose a high metabolic cost on an animal, and in that sense can be viewed as being harmful (Simpson and Raubenheimer, 2000). Other chemicals may be necessary nutrients at low concentrations but have more direct toxic effects at higher concentrations. In this model therefore different required chemicals in a food (nutrients) contribute to an overall phagostimulatory drive, which is derived from both the concentration and blend of the chemicals in the stimulus. Figure 2 shows a graphical representation of how the phagostimulatory power, shown on the vertical axis, of a chemical stimulus varies with increasing concentration and in various blends of two nutrient chemicals, shown on the horizontal axes. The phagostimulatory drive of either chemical on its own is minimal and does not show a strong dose–response relationship with concentration (the edges of the graph), but the response to each of these chemicals increases sharply when present in a blend with the other nutrient. The maximum phagostimulatory power, at the apex of the graph, coincides with the optimal blend and concentration of nutrients that corresponds to the intake target. The phagostimulatory power could be a simple sum of action potentials in chemosensory neurones reaching the central nervous system, integrating either temporally as a firing frequency, or spatially in terms of the number of phagostimulatory sensory neurones in a sensory array that respond to the stimulus (Blaney, 1975; Varanka, 1981). The non-linear response to the blends could be generated at least partially by interactions within and between sensory neurones in the periphery, examples of which have been described in many insect species (see below) and also integrated within the central nervous system
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FIG. 2 Representation of the taste model of Simpson and Raubenheimer (1996). The graph represents the relative phagostimulatory power (vertical axis) of different concentrations of two distinct chemical stimuli (horizontal axes), both singly and as blends. Neither chemical is very phagostimulatory on its own, as shown by the shallow dose–response relationship at each of the horizontal axes. The phagostimulatory power, however, increases in a non-linear way with blends of the two chemicals. The peak of the curve represents the optimal balance and concentration of the two nutrients.
(see Section 5 for a discussion). Partial supporting evidence for this model comes from studies on locusts (Simpson et al., 1991) and caterpillars of Spodoptera littoralis (Simmonds et al., 1992). In these studies chemosensory neurones of insects previously fed on balanced foods did not exhibit any strong dose–response relationship to increasing concentrations of carbohydrate or amino acids. This changes, however, when insects are pre-treated on nutritionally unbalanced foods, leaving them deficient in either carbohydrate or protein. Chemosensory neurones on the mouthparts then exhibit a selective and strong dose–response relationship specific to the nutrient in which the insect is deficient. Not only is the sensory sensitivity to the deficient nutrient increased overall, it is directed towards a particular concentration of that nutrient, such that a distinct peak-firing rate now occurs at a particular concentration that falls away at higher concentrations. This changed response profile is congruent with the peak of the phagostimulatory power shifting so that it lies over, or nearer the axis of the deficient nutrient, with the consequence that a distinct maximum response concentration can now be seen when chemically stimulated with a single chemical. Other peaked dose– response relationships have been reported in a study looking at the interactions
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between nutrient and secondary plant compounds, again in the locust (Chapman et al., 1991). There is an interaction between sucrose concentration and concentrations of various deterrent plant compounds such that there was a peak-firing rate of a sucrose-sensitive neurone for a particular mixture concentration that fell away at higher concentrations. Although there has not yet been a systematic testing of the Simpson and Raubenheimer (1996) model, formulizations such as their model provide cogent sets of predictions against which ideas about chemosensory coding can be tested and refined. The peripheral sensory physiology of chemosensation has been investigated with different emphases by different researchers. One abiding interest has been how the narrow and specialized host-plant ranges of many insects may be determined by chemosensory systems, and how changes in chemosensory responsiveness in closely related species may reflect evolutionary changes in host-plant preference (van Loon, 1990; Feeny, 1991; Roessingh et al., 1991; Mitchell and McCashin, 1994). Other research has concentrated on the more immediately adaptive use of contact-chemosensory information in the regulation of feeding and how insects use this information in conjunction with other sensory inputs and feedbacks in order to attain a balanced diet (Simpson et al., 1988, 1991; Simmonds et al., 1991, 1992; Simpson and Raubenheimer, 1996). However, these are just two different aspects of the same sensory system. Even an insect with the narrowest host-range will not be living off a homogeneous substrate; nutrients may be present in different concentrations in different parts of the plant or between plants of the same species (Schoonhoven, 1996). Conversely, a fully nutritionally adequate diet may be left untouched if it does not contain appropriate stimulatory non-food compounds or contains chemicals that are highly deterrent. Deterrent compounds are not necessarily harmful to the insect, but nevertheless their presence serves to reduce the likelihood of the insect commencing feeding on a particular food (Bernays and Chapman, 1994). Individual compounds need not be intrinsically phagostimulatory or deterrent; their perceived quality may vary with concentration, with their ratio to other chemicals present, and with the previous experience of the insect (Simpson and Raubenheimer, 1996). Relevant kinds of previous experiences may come from early life when preferences for a particular host-plant are induced and subsequently fixed (Bernays and Weiss, 1996), or with more immediate previous nutritional experience and the current nutritional needs of the insect. Chemosensory neurones frequently respond in a non-additive way to mixtures of compounds compared to when they are presented individually. Secondary plant compounds, as well as causing activity in a particular neurone, may suppress the activity of other neurones within a sensillum to other substances present in the stimulatory mixture (Mitchell, 1987; Dethier and Bowdan, 1989; Chapman et al., 1991). Suppression may also work in the other direction; activity in a salt/sugar responsive neurone in the locust Schistocerca
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americana appears to be able to inhibit spiking in a nicotine hydrogen tartrateresponsive neurone (White et al., 1990). Some compounds may simply reduce the level of activity of all the neurones in a sensillum, decreasing the amount of information reaching the CNS and phagostimulatory chemicals may amplify the response to other chemicals (Ascoli-Christensen et al., 1990). Clearly an insect sampling real plant material, containing potentially hundreds of different chemicals, may be receiving a quite different array of sensory inputs than can be guessed at from tip-recording with solutions containing perhaps at most two or three separate compounds. 2.4
VARIATION IN CHEMOSENSORY RESPONSE
The contact-chemosensory system of insects does much more than simply classify different chemicals or food types, it has an active role in shaping meal duration, the gaps between successive meals and in regulating the types of food eaten. A key method of achieving these functions is through variability in chemoreceptor response. This variability may be short term, within or between individual meals, through to longer-term variation that affects feeding strategies over course of days or over the whole life cycle of the insect. Furthermore, these longer-term changes may be plastic and reversible, or become fixed stages in post-embryonic developments. An obvious and important component of chemosensory plasticity is the adaptation of chemosensory neurones on sustained stimulation. This process can occur extremely rapidly, and phaso-tonic firing patterns are perhaps the most characteristic response of chemosensory neurones, with firing frequency decreasing rapidly within approximately 500 ms of stimulation, and may be ceasing to respond entirely after several seconds. Behaviours such as palpation (Blaney and Chapman, 1970; Mordue, 1979) and tarsal drumming may function to decrease the rate of adaptation by rapidly removing and reapplying chemoreceptors onto a stimulus. Even with such mechanisms, adaptation may have a role in regulating meal size (Bernays and Simpson, 1982), though initially its effects may be offset by the activation of a central excitatory state (Dethier, 1976) that promotes the continuation of feeding in insects once it has started regardless of continuous contact-chemosensory input (Bernays and Simpson, 1982). There are other more active mechanisms affecting chemoreceptors that may regulate feeding behaviour. The first of the shorter-term effects to be discovered was an overall depression in the responsiveness of the terminal palp sensilla of locusts immediately after feeding that slowly increases again over the course of 1–2 h and therefore appears to be distinct from more general chemosensory adaptation, which recovers more quickly (Bernays et al., 1972; Bernays and Chapman, 1972). This effect is accompanied by a general reduction in electrical conductance across the sensillum, which may arise from the physical closure of the sensillum pore, and appears to be regulated by a
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circulating hormone. Any mechanism that reduces the amount of chemosensory excitatory drive reaching the CNS will potentially have a role in determining the duration of feeding, but the population responses of contactchemosensory neurones are also subject to more specific variation. In locusts fed synthetic foods containing a surfeit of protein, the responsiveness of neurones in the maxillary palp-dome sensilla to amino acids specifically decreases (Abisgold and Simpson, 1988; Simpson et al., 1991). Similarly the responsiveness to sugars is depressed when locusts are fed foods with high levels of digestible carbohydrates (Simpson et al., 1991). The decrease in response to amino acids could be induced by injecting a solution of amino acids into the haemolymph of a ligatured maxillary palp, strongly suggesting that the effect was mediated at the level of the sensory receptors and does not require a centrifugal neural or hormonal control (Simpson and Simpson, 1992). These differences in sensory neurone responsiveness mirror the behavioural choices of a locust as it compensates for variation in the amounts of protein and carbohydrate in its diet; so that, for example, a locust previously conditioned on a high protein diet, when subsequently presented with high protein food, is far more likely to reject it, or take short meals only. Similar nutrient-specific modulation of chemosensory response has a wide occurrence in the insects and has been reported in both caterpillars (Simmonds et al., 1992) and flies (Amakawa, 2001), but may not be a feature of all contactchemosensory neurones; for example, chemosensory neurones on the legs of locusts do not vary in response according to nutritional status unlike those on the palps (Simpson, 1990). Long-term modulation of chemosensory response dependent on feeding experience may also affect chemoreceptors sensitive to non-nutrient chemicals. Exposure to foods containing aversive, but not harmful chemicals may lead to a long-lasting and specific decrease in sensitivity of contact-chemosensory neurones sensitive to those chemicals (Glendinning et al., 1999). A third category of chemosensory variation occurs over the longer-term and includes changes in responsiveness to macronutrients over the course of a day, stadium or longer, and may match the differing nutritional requirements of an insect over time, which may be dynamic in response to individual experience (Simpson and Raubenheimer, 2000) or as an intrinsic part of normal larval development (Simmonds et al., 1991; Simpson et al., 1990). Other longer term and irreversible changes in the chemosensory system may occur during post-embryonic development, ranging from induced host plant recognition, with accompanying changes in chemosensory physiology through to changes in the number of chemosensory sensilla depending on diet. Induced host recognition is a feature of many insect species, particularly caterpillars (Bernays and Weiss, 1996). In extreme cases, newly hatched caterpillars will readily accept a wide range of different foods, on which it can be reared successfully to adulthood. On first feeding on a natural host plant however, they become dietary specialists and will subsequently reject all other food
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sources, even until the point of starving to death (Szentesi and Jermy, 1990). In a recent study del Campo et al. (2001) found that induced host plant recognition in the moth Manduca sexta was accompanied by changes in the responses of chemosensory neurones in galeal sensilla, which must remain intact for host plant discrimination to occur. In particular, the responses to a characteristic host secondary plant compound increased strongly relative to the phagostimulant glucose. Nutritional experience may have a strong effect on the number of chemosensory sensilla expressed during post-embryonic development in insect species in which the number of sensilla increases with every moult, such as the orthopteroid groups (Chapman and Lee, 1991; Rogers and Simpson, 1997; Bernays and Chapman, 1998). Grasshoppers reared in an impoverished chemical environment develop fewer chemosensory sensilla than those reared in environments with a greater variety of chemical stimuli. In these experiments an impoverished chemical environment was usually provided by rearing the insects on completely nutritionally balanced but otherwise chemically bland synthetic foods. Increasing chemical diversity through allowing the insects access to different foods, containing either different non-nutrient flavours (Rogers and Simpson, 1997; Bernays and Chapman, 1998) or varying in their nutritional balance (Rogers and Simpson, 1997) increased sensilla formation. Forcing a locust to regulate nutrient intake through assessing and choosing between nutritionally unbalanced but complementary foods increased sensilla formation compared to single balanced foods, even though the total number of different chemicals present was exactly the same in both situations (Rogers and Simpson, 1997). The effect appears to be locally driven; locusts fed bland synthetic foods in the presence of plant odours expressed increased numbers of olfactory sensilla on the antennae, but the numbers of gustatory sensilla on the palps were lower than locusts actually fed on plant material (Rogers and Simpson, 1997). All of these effects indicate that the contact-chemosensory system has a dynamic role in the regulation of feeding and food choice and its tasks are much more varied than that of passive recognition, or simply discriminating between acceptable and unacceptable food types. Modulation of olfactory sensitivity is also known to occur in insects (Bowen et al., 1988; den Otter et al., 1991), and inhibition of olfactory sensory neurone responsiveness after feeding also may have a role in the regulation of meals (Takken et al., 2001). In mosquitoes the decrease in olfactory sensitivity following blood feeding is accompanied by decreased expression of a particular candidate olfactory receptor protein (Fox et al., 2002). These processes taken together are perhaps representative of the observation that invertebrate nervous systems do more at the periphery or in the early stages of integration than vertebrate nervous systems where various kinds of central nervous modulation of taste inputs have been described (Jacobs et al., 1988; Rolls, 1989).
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Central projections of sensory neurones
Determining the destination and branching patterns of sensory neurones within the central nervous system has made a major contribution to our understanding of how signals from several different sensory modalities are processed. Sensory neurones and the interneurones that receive input from different modalities commonly have branches in specific regions of insect ganglia, indicating that different sensory modalities are represented and processed in specific areas. For example, proprioceptive sensory neurones from receptors that monitor movements about limb joints have branches of fine neurites predominantly in dorso-lateral regions of the ganglion (the Lateral Association Centres) (Burrows, 1987; Pflu¨ger et al., 1988) while sensory neurones from exteroceptive tactile hairs project into a ventral region of the ganglion known as the Ventral Association Centre (VAC) (Newland, 1991; Pflu¨ger et al., 1988). As well as segregating into different modality-specific processing regions within the central nervous system, another major feature of sensory systems is that the projections of their sensory neurones form orderly maps within these modality-specific regions. These orderly projections may form consistent and predictable central representations of the location of sensory receptors on or in the body, i.e. a somatosensory map, or they may be organized according to particular coding properties of the sensory neurones themselves. The central projections of hairs on the limbs of invertebrates (Murphey et al., 1980; Murphey, 1981; Pflu¨ger et al., 1981; Johnson and Murphey, 1985; Levine et al., 1985; Kent and Levine, 1988; Peterson and Weeks, 1988; Newland, 1991) and vertebrates (Brown et al., 1977, 1980) have been shown to form somatotopic maps in such a way that the spatial location of a receptor on the limb is preserved within the map. Conversely, the central projections of sensory neurones from olfactory receptors on the antennae form odotopic maps in which sensory neurones segregate into compartments, or glomeruli, within the olfactory neuropil of the brain (Hildebrand and Shepherd, 1997). In the auditory system, sensory neurones form a tonotopic map (Oldfield, 1982; Ro¨mer, 1983; Ro¨mer et al., 1988) in which sensory neurones project to particular regions of the nervous system, depending on the sound frequency they respond to best, and sensory neurones from the eyes are arranged retinotopically (Strausfeld, 1976). In these latter two cases the maps combine features of both somatosensory and coding property organization. Such sensory mapping studies have generated an expectation of how contact-chemosensory neurones could or even should be organized within the central nervous system. Unfortunately though, far fewer studies have been directed towards understanding the organization of sensory neurones innervating taste receptors, and taken together they make a rather confusing and sometimes contradictory pattern. As mentioned previously, the contactchemosensory sensilla of insects frequently cover the mouthparts, body and
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limbs (Chapman, 1982) and are often multimodal in that they contain one mechanosensory neurone and several chemosensory neurones (Blaney and Chapman, 1969; Kendall, 1970; Dethier, 1976; Klein, 1981; Murphey et al., 1989b). These features, combined with the small diameter of chemosensory axons – often less than 1 mm – makes the staining of their terminal arborizations very difficult, especially given the large distances they project from the periphery to the central nervous system. Most studies have utilized a retrograde dye filling technique using cobalt chloride (Pitman et al., 1972) with subsequent silver intensification (Bacon and Altman, 1977). Even with large diameter axons this method of staining can have low success rates over large distances (see for example Newland, 1991). The more recent development of the neurobiotin retrograde filling technique for use in insects (see for example Consoulas et al., 1996) has provided a far better means of staining small diameter neurones and processes over large distances – up to 4–5 cm for receptors on insect limbs (Tousson and Hustert, 2000). These dyes are introduced into the sensory neurones by cutting a sensillum and severing the sensory dendrites contained within it, allowing access by the dye, which is then carried down the axons of the neurones to their final arborizations in the central nervous system. This method does not usually allow any targeted selection of one sensory neurone type over another although some workers have reported success with selective staining using a method in which tastants (Shanbhag and Singh, 1992) are incorporated in the stain solution applied to cut sensilla. How this works in cut neurones, and whether other biochemical processes rather than selective neural activation is responsible for the selective staining, is not clear. Assigning modality or chemosensory sensitivity to the multiple axons stained from an individual chemosensory sensillum is still largely reliant on inference. Nevertheless, a number of recent studies have revealed how sensory afferents from bimodal sensilla are organized in the central nervous system and how these are related to the projections of sensory neurones from tactile hairs. In doing so, these studies have indicated the likely postsynaptic targets and possible function of the taste receptors in one species of insect, the locust. 3.1
CENTRAL ORGANIZATION OF SENSORY NEURONES FROM TACTILE HAIRS
Bimodal chemo/mechanosensory sensilla (Fig. 3A–C) are intermingled with unimodal tactile hairs containing only a single mechanosensory neurone on the surfaces of many insects (Fig. 3D–F). It might be expected that the mechanosensory neurones from bimodal sensilla would follow a similar organization to the mechanosensory neurones from tactile hairs in the central nervous system, since both encode similar information (Newland and Burrows, 1994) and both contribute to the receptive field properties of mechanosensory interneurones that receive convergent synaptic inputs from both types of mechanosensory neurones (Burrows and Newland, 1994). Understanding the
FIG. 3 Innervation of exteroceptors on the locust leg. (A–C) Contact chemoreceptors (basiconic sensilla) are bimodal and multiply innervated. (A) Electron micrograph of a basiconic sensillum on the hind leg of the locust. (B) Backfilling a basiconic sensillum on the tarsus results in the staining of five sensory neurones, four of which are chemosensitive and one mechanosensitive. (C) Placing an electrode filled with 250 mM sodium chloride over a basiconic sensillum evokes a burst of action potentials of different amplitudes (chemosensory responses). Displacement of the shaft of the sensillum (arrow right side) evokes a burst of action potentials with larger amplitudes from the mechanosensory neurones. (D–F) Tactile hairs (trichoid sensilla) are singly innervated. (D) Electron micrograph of long tactile hairs on the hind leg of the locust. (E) Retrograde filling of a cut tactile hair on the dorsal tarsus of the mesothoracic leg with neurobiotin results in the staining of a single sensory neurone in the mesothoracic ganglion. (F) Movement of an electrode placed over the cut shaft of a tactile hair, filled with 50 mM sodium chloride, deflects the hair (arrows) and evokes bursts of action potentials of a single amplitude in its sensory neurone. Based on Newland et al. (2000), Journal of Comparative Neurology ß 2000 Wiley-Liss, Inc.
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organization of sensory neurones from tactile hairs in the central nervous system should therefore allow a putative separation of mechanosensory from chemosensory neurones in projections from bimodal sensilla. Fortunately, we already know a great deal about how the individual sensory neurones from exteroceptive tactile hairs on the legs project within the central nervous system. All sensory neurones from mechanosensory sensilla on the leg project within their local segmental ganglion or neuromere. The arborizations of these sensory neurones form a complete 3-dimensional somatotopic map of each segment of the leg, and of parts of the thorax in the VAC (Newland, 1991; Mu¨cke and Lakes-Harlan, 1995; Newland et al., 2000). Taking the middle leg of the locust as an example, the long axis of the leg is represented by a region running along a medio-lateral axis within the mesothoracic ganglion (Fig. 4A–C, shaded region) (Newland et al., 2000). Within this map, sensory neurones from femoral hairs project most medially and slightly anteriorally (Fig. 4A), sensory neurones from tibial hairs arborize more centrally (Fig. 4B) and sensory neurones from tarsal hairs project posterio-laterally towards the edge of the neuropil (Fig. 4C). Similarly, sensory neurones from hairs located dorsally on the leg project more dorsally in the VAC than those located ventrally on the leg, and hairs located on the anterior surface of the leg project more anteriorally than those located on the posterior surface (Newland, 1991; Newland et al., 2000). A fundamentally similar organization is seen in the metathoracic ganglion, which serves the hind legs (Newland, 1991). These somatotopic maps follow a well established and conserved pattern also shown for leg bristles afferents in other insects such as in crickets (Johnson and Murphey, 1985) and flies (Murphey et al., 1989b). 3.2
CENTRAL ORGANIZATION OF SENSORY NEURONES FROM CONTACT-CHEMOSENSORY SENSILLA
Given that other modalities of sensory neurones show considerable segregation within the central nervous system (Burrows, 1987; Pflu¨ger et al., 1988), it is possible to make a number of predictions about the projections of bimodal contact-chemosensory sensilla, and knowing the projection regions of mechanosensory neurones from tactile hairs it should be possible to separate mechanosensory from chemosensory processing regions. On the basis that general organizational principles apply one would predict: First, that the chemosensory and mechanosensory afferents from contactchemosensory neurones would show spatial segregation. Second, the mechanosensory afferents of the contact-chemosensory sensilla should project to similar regions of the central nervous system as tactile hair afferents.
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FIG. 4 Mapping of sensory neurones from tactile hairs on the proximo-distal mesothoracic leg axis. (A) The central projection of a sensory neurone from a tactile hair on the dorsal surface of the femur. (B) The central projection of a sensory neurone from a hair on the dorsal tibia. (C) The central projection of a sensory neurone from a hair on the tarsus. The stippled area indicates the area occupied by the branches of the sensory neurones showed in A–C. Inset shows the locations of the tactile hairs on the mesothoracic leg. Based on Newland et al., 2000, Journal of Comparative Neurology ß 2000 Wiley-Liss, Inc.
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Third, the mechanosensory afferents of the contact-chemosensory sensilla would be arranged somatotopically. Fourth, that different chemosensory neurones from within a single receptor may show segregated projections based on the quality of chemical they respond to best, by analogy with the odotopic organization of the antennal lobe. Each of these predictions is addressed in turn below. 3.3
MODALITY-SPECIFIC SEGREGATION
Based on available evidence, there seem to be considerable differences between the Diptera and other insect groups in the organization of the central projections of bimodal sensilla. Studies on the flies Phormia and Drosophila (Edgecomb and Murdock, 1992; Shanbhag and Singh, 1992; Murphey et al., 1989a) have suggested that there are modality-specific projections from bimodal sensilla on the labellum and tarsi. Cobalt chloride dye filling results in a number of stained sensory neurones, one of which is usually of a thicker diameter than the others and projects to a region of neuropil occupied by afferents from mechanosensory bristles, and is hence thought to be mechanosensitive (Murphey et al., 1989a). The remaining finer diameter processes that project to a more ventro-medial region of neuropil are presumed to be chemosensory. Similar observations have been made by Edgecomb and Murdock (1992) from taste receptors on the labellum of Phormia, providing good evidence of spatial segregation of different modalities in these insects. Our recent studies analysing sensory projections from bimodal sensilla on the middle leg of the locust, by contrast show no consistent differences in the morphology of particular sensory neurones from a given basiconic sensillum (Newland et al., 2000). There were no consistent differences in the diameters of stained neurones and all sensory neurones from a single receptor projected into the ventral association centre (Fig. 5A–C) in which we know the tactile afferents arborize, and there appeared to be no dorsoventral segregation of different fibres from the same sensillum (Newland et al., 2000). Tousson and Hustert (2000) recently undertook a detailed analysis of contact-chemosensory sensilla on the ovipositor valves that are moved rhythmically in opening and closing movements as the abdomen is driven through the sand during egg-laying (Thompson, 1986a,b). The chemoreceptors, distributed over both pairs of valves, provide information about the chemical composition of the sand or soil in which the locust lays its eggs and have a major influence on egg-laying behaviour (Woodrow, 1965). Tousson and Hustert (2000) found neither consistent differences in axon diameters that could be attributable to the modality of a sensory neurone nor a spatial segregation of the sensory neurones from a given receptor. Likewise
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FIG. 5 Sensory neurones from contact-chemoreceptor sensilla along the proximodistal leg axis. (A) An example of the central projection of sensory neurones from a basiconic sensillum located on the proximo-dorsal femur. (B) Projection of sensory neurones from a basiconic sensillum on the proximo-dorsal tibia. (C) Central projection from a basiconic sensillum located on the dorsal tarsus. The light stippling represents the area occupied by the sensory neurones from all the basiconic sensilla. The darker stippling indicates the projection areas of tactile hair afferents from similar proximodistal locations of the middle leg taken from Fig. 4. Not only do the sensory neurones from the basiconic sensilla project to regions specific for particular regions of the leg but there is also a close correlation between the branching areas of sensory neurones from both classes of receptor. Based on Newland et al. (2000), Journal of Comparative Neurology ß 2000 Wiley-Liss, Inc.
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FIG. 6 Projections of sensory neurones from two adjacent basiconic sensilla on the tip of a maxillary palp of a locust, which play an important role in food sampling by the mouthparts. (A) Projections in the sub-oesophageal ganglion. A total of 14 sensory neurones enter the ganglion through the maxillary nerve (i) and three axons (iv) ascend to the brain through the circum-oesophageal connectives (top), but within the suboesphageal ganglion (ii) there is no clear segregation of the projections from the different modalities of sensory neurones in whole mount or in section. Neurites from three sensory neurones project back towards the labial neuromere (iii) where they overlap with an equivalent forward projection from labial palp sensilla. Anterior is to the top (B–E). Transverse 45 mm sections through the suboesophageal ganglion showing the sensory projections at the levels indicated in (A). Based on Rogers (1998).
Rogers (1998) using cobalt back-filling techniques traced the sensory projections of contact-chemosensory sensilla on the tips of the maxillary and labial palps of locusts and found no differences in axon diameters between the sensory neurones from a given receptor. In no instances could a single, presumably mechanosensory, neurone be shown to arborize in a region spatially distinct from the branches of all the other stained neurones (Fig. 6). In summary, in the locust both mechano- and chemo-sensory neurones from bimodal sensilla project to the same region of neuropil in the ventral association centres as sensory neurones innervating tactile hairs (Newland
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et al., 2000; Tousson and Hustert, 2000). The branches of all the sensory neurones from a given sensillum are intertwined. All basiconic sensillum afferents branch within a restricted dorso-ventral region within the ventral association centre regardless of the route the axons take to reach their arborization site (Figs. 3E, 5A–C). Indeed, sensory axons travelling in the peripheral nerve and across the ganglion can take quite divergent paths before converging on their final arborization region. Kent and Hildebrand (1987) stained sensory projections from mouthpart structures of the larval moth, Manduca sexta, and similarly found no obvious modality-specific segregation; there was no indication that a minority of neurones from each sensillum arborized in regions spatially distinct from those of other neurones: though in this study projections from small groups of sensilla rather than individuals were stained for the most part, perhaps making interpretation more difficult. Given the conserved nature of the organization of sensory neurones from tactile hairs/bristles across many species it is somewhat surprising to find that the organization of sensory neurones from contact-chemosensory sensilla varies between insect orders, although being consistent within a single species. The detailed analyses of the central projections of locust contact-chemosensory sensilla on legs, ovipositor and mouthparts – structures that are involved in several different behaviours – all appear to show similar organizational features. Sensory projection patterns in Manduca, appear to be more similar to locusts than those of the Diptera. Why the locust and Manduca should differ so markedly, but consistently, from the Diptera is unclear but it may imply that contact-chemosensory information may serve a different role in these two groups of insects.
3.4
SOMATOTOPIC MAPPING OF SENSORY NEURONES INNERVATING CONTACT-CHEMOSENSORY SENSILLA
In locusts, the most striking feature of the central projections from bimodal sensilla is that all the sensory neurones, both mechanosensory and chemosensory, form a somatosensory map in which the spatial position of the contact-chemosensory sensillum on the periphery is preserved in the central nervous system (Newland et al., 2000). Our results have clearly shown that chemosensory neurones form somatotopic maps in the same way as tactile afferents (Newland et al., 2000). Given the convergence of mechanosensory inputs from both tactile hairs and bimodal sensilla onto the same local interneurones (Burrows and Newland, 1993, 1994), it would have been surprising if the mechanosensory afferents from contact-chemosensory sensilla had not followed a similar somatosensory organization as tactile hairs. What is perhaps more surprising is that all the sensory neurones from contactchemosensory sensilla appear to follow a purely somatosensory organization,
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an organization that has important implications for the processing of chemosensory information, as detailed below (Sections 4 and 5). On the labellum of the blowfly there are 11 identifiable long contactchemosensory sensilla, each innervated by a single mechanosensitive afferent and four chemosensory afferents, each with different chemical sensitivities (Dethier, 1976). Yetman and Pollack (1986) found that the mechanosensitive afferents from these contact-chemosensory sensilla formed a discontinuous map that reflected the spatial position of the receptor on the labellum, a finding supported by Edgecomb and Murdock (1992). In an analysis of the central projections of contact-chemosensory sensilla on the fore leg tarsus, Murphey et al. (1989a) suggest that there is a spatial segregation of chemo- and mechano-sensory neurones from contact-chemosensilla. While that study did not specifically address the problem of the topographic mapping, it did however show that the putative mechanosensory neurone from contactchemoreceptors projected to a region in the ganglion that receives inputs from tarsal mechanosensory bristle neurones (Murphey et al., 1989b). The putative chemosensory neurones from the same receptors, however, project more medially in the central nervous system. There is a potential problem here, as the chemosensory afferents from tarsal chemosensilla appear to project to a region likely to be occupied by mechanosensory neurones from tactile bristles on more proximal leg segments (Murphey et al., 1989b). Further studies may help to resolve this apparent paradox. What is now needed is to extend these analyses to determine whether there is a precise topographic map in all insects. These few studies therefore appear to indicate a general principle that the mechanosensory afferents from contactchemosensory sensilla are organized somatotopically, and project to the same region as sensory neurones form purely exteroceptive sensilla. However, there appears to be a divergence between groups of insects in which projections of mechanosensory and chemosensory neurones are segregated. 3.5
CHEMOSENSORY MAPPING
In addition to modality-specific segregation, a further important question is whether there is any evidence for the spatial segregation of chemosensory afferents with different chemical sensitivities, i.e. is there any evidence for chemotopic mapping? In separate studies on the locust leg (Newland et al., 2000), and ovipositor (Tousson and Hustert, 2000) there was no evidence for the segregation of neurones that could in any way be based on chemical quality. Instead, as we have already seen, all neurones ramified in the same area of sensory neuropil, and branches from different neurones were largely intertwined. Projections from contact-chemosensory sensilla on the maxillary and labial palps of locusts have a somewhat more complex organization (Rogers, 1998). Most neurones from sensilla on the maxillary and labial palp dome arborized within the ventral neuropil of their respective neuromeres, and
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resemble the arborizations seen in projections from leg contact-chemosensory sensilla, with no obvious spatial segregation of different neurones. The arborizations of neurones from individual palp dome sensilla, however, occupied a far larger volume of this neuropil compared with projections from individual leg bimodal sensilla, stretching from the lateral edge of the neuropil to near the midline. This presumably reflects the need to accommodate several thousand palp dome sensory neurones (Chapman, 1982) in the same space. The apparent dominance of chemosensory afferents in the sub-oesophageal ganglion may reflect the importance of these inputs to the central nervous system from the mouthparts, in which food assessment is a far more central role than on the legs, with the palp-dome sensilla being the largest source of sensory neurones from the maxillary palp (Blaney and Chapman, 1969). As sensory projections from different individual palp-dome sensilla all occupied the same extent of space within the maxillary or labial neuromeres, cutting and staining the afferents from an entire palp-dome led to a much denser, but no more extensive, region of staining in the sub-oesophageal ganglion. Branches from at least two neurones per sensillum occupied a longitudinal-medial region of the sub-oesophageal ganglion, with overlapping projections from both maxillary and labial palp-dome sensilla. Although most neurones from palp-dome sensilla branched entirely locally within the sub-oesophageal ganglion, in approximately 30% of successful stains 1–3 neurones per sensillum ascended to the brain and terminated in a region, the lobus glomerulatus, known from histological studies to have a fine glomerular structure (Ernst et al., 1977). These ascending neurones appeared not to branch at all in the region of the sub-oesophageal ganglion where the other neurones arborized, and provide an exception to the apparent nonsegregation of chemosensory afferents from contact-chemosensilla in locusts. The proportion of projections containing ascending axons was far greater than would be expected if projections from only the multiporous olfactory sensilla on the palp dome (Blaney, 1977) ascended. Even in those projections containing ascending axons, however, it was only ever a small proportion of the total number of neurones that travelled to the brain, which suggests that most processing of chemosensory information from the mouthparts must be carried out locally within the sub-oesophageal ganglion. Ascending projections from neurones in purely mechanosensory sensilla have been described (Bra¨unig et al., 1981), but typically these are only found in projections from sensilla near the midline of the body, in contrast to the palp-dome sensilla. Allowing for the relative developmental distortion of the sub-oesophageal ganglion in different orders of insects, a broadly similar pattern of projection has been reported, if not from individual palp-bimodal sensilla, but from the homologous nerves or palps in a number of different insects. For example, the projections from the maxillary and labial palps of Manduca sexta (Kent and Hildebrand, 1987) and from the whole maxillary palp dome of the cockroach
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Supella longipalpa (Prakash et al., 1995) show a great deal of similarity with those of locusts. These results contrast with the organization of olfactory afferents of insects in which there is an odotopic map (Vickers et al., 1998) in which many glomeruli in the deuterocerebrum of the antennal lobe receive input from different functional populations of olfactory afferents. In this way a given odour is encoded in the antennal lobe by the summed activity of specific, but overlapping, combinations of glomeruli. The apparent lack of such a gustatory map in the locust may again point to a different function of the taste receptors on the legs and perhaps the mouthparts as well. In Phormia and Drosophila there is evidence to suggest a segregation of several, although not all, of the chemosensory neurones from labellar bimodal sensilla, again contrasting with locusts. Individual chemosensory neurones send clearly spatially distinct branches to one, or more than one, of four distinct regions in the sub-oesophageal ganglion (Yetman and Pollack, 1986; Murphey et al., 1989b; Edgecomb and Murdock, 1992). Shanbhag and Singh (1992), finding a similar arrangement in Drosophila, have interpreted it as glomerular and analogous to the antennal lobes, but their total number of proposed glomeruli is only seven across the entire sub-oesophageal ganglion. It appears that contact-chemosensory neurones in flies are coarsely compartmentalized in the central nervous system, unlike those from olfactory sensilla. Earlier in this section we hypothesized about the organization of the sensory afferents from the bimodal basiconic sensilla. As we predicted, the mechanosensory afferents of taste receptors project to similar regions of the central nervous system as tactile hair afferents and exhibit a somatosensory organization. Contrary to what we might have expected, however, only in some insect species do chemosensory and mechanosensory afferents show a spatial segregation and different chemosensory neurones from within a single sensillum show segregated projections based on chemical sensitivity. The organization found in the locust is similar in some respects to that found in vertebrates where sensory neurones in the cranial nerve innervating the taste buds form parallel maps in the brain together with mechanosensitive neurones in the trigeminal nerve (Spector, 2000). It appears likely that the behavioural function of this is to allow the localization of food particles in the mouth. Similarly, for the locust the parallel organization of taste and touch also provides information about the spatial location of gustatory stimuli on a leg.
4
Local circuits and their role in processing gustatory signals
Until recently little was known about the central processing of gustatory information in any insect. Studies by Mitchell and Itagaki (1992) and Rogers
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and Simpson (1999) described responses of central neurones in the suboesophageal ganglion to taste stimuli in the blowfly Phormia and the locust, Locusta migratoria, respectively. Although neurones were found exhibiting differential responses to different chemical solutions or leaf types, they could not be placed in a wider context of contact-chemosensory processing. Therefore the role of these neurones in chemosensory information coding, categorization, further processing and use in cueing appropriate behavioural responses could not be systematically assessed. Dethier (1973, 1976), Schoonhoven (1987) and Schoonhoven and Blom (1988) have proposed theoretical models of central taste processing, by extrapolating from recordings of sensory neurones at the periphery and feeding behaviour. These models posit that chemosensory signals, either directly from different chemosensory neurones in a population of sensilla or from the output of an across-fibre patterning array, are summed algebraically by the central nervous system. Thus, some chemical stimuli are coded positive (phagostimulants), others from deterrent-detecting neurones coded negative and these inputs are simply added by the central nervous system. If the total positive drive, expressed in terms of the firing frequency of all phagostimulatory-coding neurones is greater than the negative drive from all the deterrence-coding neurones then feeding will occur, and furthermore the amount of food eaten is linearly correlated with the net positive drive. In caterpillars there appears to be a strong correlation between chemosensory neurone firing rates and feeding performance if gustatory stimuli are presented singly in otherwise inert non-nutritious foods (Blom, 1978; Schoonhoven and Blom, 1988), but such clear correlations break down if more complex, chemically varied foods are fed (de Boer, 1993). If food selection was simply made according to such additive models then food selection in insects in which some chemosensilla have been unilaterally removed (thus decreasing overall sensory input) would be expected to be severely impaired, which does not appear to be the case (de Boer, 1993). Moreover, quite different chemical stimuli put together in blends could potentially provide the same overall or net phagostimulatory drive depending on how the central nervous added or subtracted the contribution of each chemical in the blend, possibly leading to very broad range of acceptable foods, not all of which may provide adequate nutrition or protect against the ingestion of harmful substances. For example, a harmful or deterrent chemical in a blend could be entirely negated by a superabundance of a phagostimulant, regardless of its intrinsic unsuitability. Recently we have made what we believe is significant progress in understanding the central processing of chemosensory information in one particular experimental system, that of the metathoracic ganglion in the locust. This may not seem like the most obvious choice for investigating central gustatory processing in an insect, but as we detail below, it offers several important advantages as a model system.
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To understand gustation at a central level, it is vitally important to know something of the neurones and pathways through which information flows in the central nervous system in order to give a context to recordings from central neurones. Here the locust as a model nervous system comes into its own, since we have extensive knowledge of the organization of its nervous system, and in particular the local circuits in the metathoracic ganglion that receive sensory information from, and control movements of, the hind leg (reviewed by Burrows, 1996). Sensory receptors on and in just one hind leg send some 10 000 sensory axons to the local circuits in the metathoracic ganglion (Fig. 7A). These neurones come from different modalities of receptor, including, those that detect movement about the limb joints, stresses on the cuticle, touch, and taste. It has been estimated however, that approximately 80% of all sensory neurones from limb receptors are from the contact-chemosensory sensilla (Burrows, 1996). In this section we give a brief overview of the organization of local circuits in the metathoracic ganglion and the evidence that they process chemosensory signals. A key feature of local circuits is that the reflex pathways are short, which means that there are only one or two synapses between a sensory signal and a motor response. The local circuits that control the movement of a hind leg comprise approximately 1000 interneurones, just over half of which are local interneurones (Fig. 7B and C), so called because they lack an axon, and hence have all their branches restricted to one part of the nervous system. These local interneurones are of two distinct types; those that communicate by all-or-nothing action potentials or spikes (spiking local interneurones, Fig. 7B), and those that communicate by means of graded electrical signals (non-spiking interneurones, Fig. 7C). A third class of neurone, the intersegmental interneurone (Fig. 7D), has either an ascending or descending axon. These axons project to other local circuits where they are involved in intersegmental coordination. Finally, there are approximately 80 motor neurones that activate the muscles of a hind leg (Fig. 7E). These different neuronal types provide the architecture within which movements of the leg are produced and controlled. The mouthparts are paired appendages similar to the legs and it is very likely that similar circuits underlie the organization of their sensory processing and motor control.
4.1
PROCESSING OF SENSORY SIGNALS
Sensory signals from receptors on a leg, including contact-chemoreceptors, are processed initially by spiking local interneurones. Three distinct populations of these interneurones have so far been described in the metathoracic ganglion with homologous populations in other ganglia, which were identified initially on the basis of the location of their somata.
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FIG. 7 Local circuit neurones in the metathoracic ganglion, which are components of the networks controlling limb movements. (A) Drawing of the central projections of a sensory neurone innervating a tactile hair on the femur. (B) The dorsal and ventral branches of a spiking local interneurone with its cell body located at the ventral midline of the ganglion. A single process links the two fields of branches. (C) A non-spiking interneurone that contributes to the control of sets of leg motor neurones. (D) A drawing of an intersegmental ascending interneurone with its main branching field in the metathoracic ganglion, and (E) a motor neurone that innervates the flexor tibiae muscle of the hind leg. Reprinted from Newland and Burrows (1997) ß 1997, with permission from Elsevier Science.
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Most of what we know of the spiking local interneurones comes from extensive studies of a population with somata located at the midline of the ganglion, which release the inhibitory neurotransmitter -amino butyric acid (GABA) (reviewed by Burrows, 1996). These interneurones have been shown to be crucially involved in the processing of exteroceptive (Burrows, 1992; Burrows and Siegler, 1982, 1984; Siegler and Burrows, 1983, 1984) and proprioceptive signals (Burrows, 1987), as well as signals arising from mechanical deformation of the cuticle (Newland and Emptage, 1996), and most recently to chemical stimuli (Newland, 1999). Each interneurone responds to stimulation of a particular array of receptors on a specific region of the body or legs, which define its receptive field. For tactile signals from hairs (trichoid sensilla) on the leg, different members of the population of midline interneurones have receptive fields covering different regions of the leg, so that the entire leg is mapped across the population of interneurones (Burrows, 1992). Newland and Burrows (1994) found that the mechanosensory afferent from bimodal basiconic sensilla converge onto the same sets of spiking local interneurones that process signals from neighbouring tactile hairs and together they contribute to the receptive field of the interneurones. The shapes of both spiking local and intersegmental interneurones and the pattern of their branches within the central nervous system are related to the pattern of input and output connections that each makes with other neurones. The branching pattern of spiking local interneurones is related to the somatotopic map of the arrays of sensory afferents from which they receive inputs (Newland, 1991; Burrows and Newland, 1994). Spiking local interneurones that receive direct input from exteroceptive neurones have two fields of branches. Interneurones receive predominantly input synapses from sensory neurones in the ventral part of the metathoracic neuropil, while in more dorsal neuropil the interneurones generally make output synapses (Watson and Burrows, 1985). Crucially, the receptive fields of the spiking local interneurones are specified by the overlap of their ventral branches with a particular spatial array of sensory afferents in the somatotopic map (Figs. 4 and 5). Interneurones with extensive branches in the ventral neuropil have extensive receptive fields on the leg, while other interneurones with more restricted ventral branches have restricted receptive fields. For example, spiking local interneurones with branches in the posteriorlateral region of the ventral neuropil, where tarsal afferents project, have receptive fields confined to the tarsus, while those with branches in medialanterior neuropil, to which femoral afferents project, have receptive fields on the femur. Thus there is a clear correlation between the receptive fields of local interneurones, their neurite morphology, and the central projections of mechanosensory neurones. Original observations by Slifer (1954, 1956) suggested that contactchemoreceptors on the leg were able to detect certain odours, which subsequently led to the production of avoidance movements virtually identical to those elicited by tactile stimulation (Pflu¨ger, 1980). This led us to consider
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the role of gustatory information in the production of local reflex movements, how tastes were detected and what central neurones were responsible for the processing of such information. As shown in Section 3 above, the chemosensory neurones from particular regions of the leg project to exactly the same region of neuropil as afferents from mechanosensory receptors, and furthermore they make convergent synaptic inputs onto the same spiking local interneurones (Fig. 8A–D). At least one chemosensory neurone in each basiconic sensillum responds to the odours of weak acids, such as acetic or formic acids, directed towards the leg. These odours are able to activate chemoreceptors in the absence of any mechanosensory input, unlike more conventional gustatory stimuli, and elicit strong activity in the same spiking local interneurones that respond to touch. Given the anatomical similarity of the sensory projections synapsing onto them it might be predicted that the chemoreceptive and mechanoreceptive fields of interneurones would be similar and overlapping. This is indeed the case (Newland, 1999), with the chemosensory receptive fields of spiking local interneurones mapping the surface of a leg so that spatial information relating to the location of a taste stimulus is preserved (Fig. 8E). The receptive fields of most interneurones tested were similar for mechanosensory and chemosensory inputs. Moreover, the magnitude of response in interneurones during chemosensory stimulation varies in a graded manner along the long axis of the leg, thus creating gradients in the chemosensory receptive fields of interneurones in much the same way as occurs in mechanosensory receptive fields. We demonstrated recently that chemosensory afferents from basiconic sensilla make monosynaptic inputs onto the same midline spiking local interneurones that receive monosynaptic exteroceptive inputs (Fig. 9; Newland, 1999). Normally, applying a chemical solution, e.g. a droplet of 100 mM NaCl, to the leg evokes a stronger response in a local interneurone than that seen in response to an equivalent purely mechanosensory stimulus such as a drop of water (Fig. 9A). Individual postsynaptic potentials evoked by mechanosensory afferents are much larger (Fig. 9B) compared with those evoked by chemosensory neurones in spiking local interneurones. As well as evoking small postsynaptic potentials, the chemosensory neurones from basiconic sensilla adapt rapidly to sustained stimulation. Newland (1999) therefore used electrical stimulation of individual basiconic sensilla to demonstrate that more than one of the neurones in the sensillum make monosynaptic connections onto local interneurones. The size of post-synaptic potential in the interneurones increased stepwise as increasing applied current activated more neurones in the sensillum. An individual action potential, which is activated in an all-or-nothing manner by electrical stimulation, elicits a unitary sized post-synaptic potential; only the near-synchronous arrival of two
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FIG. 8 Receptive field properties of spiking local interneurones. The receptive fields on the leg that provide input to spiking local interneurones are similar in location for both mechano- and chemosensory inputs. (A) An interneurone with a chemosensory receptive field restricted to the dorsal tarsus received mechanosensory inputs only from the tarsus (B) and not from any other parts of the leg. (C) A different interneurone with a chemosensory receptive field restricted to the femur likewise received mechanosensory inputs only from the femur (D) and not the tarsus. (E) The receptive fields of a further four types of spiking local interneurone from the ventral midline population. To the left of each pair of diagrams is the chemosensory field and to the right is the mechanosensory field. The diagrams show the receptive fields of the interneurones as if a hind leg were opened with a ventral midline incision and laid flat (as shown in inset). Excitatory regions of a receptive field are shown crosshatched and inhibitory regions are stippled. The chemosensory receptive fields were determined using stimulation with acetic acid odour. a, anterior, p, posterior, d, dorsal and v, ventral. Based on Newland (1999).
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FIG. 9 Spiking local interneurones receive convergent chemo- and mechanosensory inputs. (A) Droplets of water or 100 mM sodium chloride applied to the hind leg evoke depolarization and spikes in an interneurone. The duration of the response is longer when sodium chloride is applied. (B) Mechanosensory spikes, evoked by deflecting a sensillum, are each followed by excitatory post-synaptic potential in an interneurone. (C) Superimposing the traces of the oscilloscope reveals that each spike is followed by an EPSP with a short and constant latency typical of a monosynaptic connection. Low levels of electric stimulation evoke EPSPs with amplitudes similar to that evoked by mechanosensory stimulation. Stimulating basiconic sensilla at suprathreshold levels (dotted lines) leads to the recruitment of a delayed depolarization that occurred with a constant latency, that sums with the potential evoked at threshold levels (solid lines), and which is presumed to be chemosensory. Based on Newland (1999).
or more action potentials, produced by passing the spike generation threshold of different neurones could increase the size of the postsynaptic potential in this way. In summary: Spiking local interneurones receive convergent monosynaptic inputs from (a) the chemoreceptors from arrays of basiconic sensilla (Newland, 1999); (b) the mechanosensory neurone innervating each of the same basiconic sensilla (Burrows and Newland, 1994; Newland and Burrows, 1994); and (c) purely tactile hairs (Burrows, 1992). The two types of sensilla (chemosensory and mechanosensory) on the leg are intermingled and provide a contiguous array of converging inputs to an interneurone.
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Within the extent of a receptive field, not all receptors contribute to the receptive field. For example, some interneurones receive input only from highthreshold tactile hairs while others receive input from low-threshold hairs (Burrows and Newland, 1994). All interneurones that receive mechanosensitive input also receive chemosensory input (Rogers and Newland, 2002). Not all receptors contribute equally to a receptive field so that gradients of excitability are produced with one area providing the strongest input. This area can be different for different interneurones. The same is also true for chemosensory inputs (Newland, 1999). The organization of the receptive field of an interneurone appears to be consistent from animal-to-animal (Burrows and Newland, 1993). Sensory neurones make divergent patterns of connections with members of different populations of spiking local interneurone, with intersegmental interneurones, non-spiking interneurones and motor neurones so that there is considerable distributed processing of sensory signals. Intersegmental interneurones receive synaptic inputs in one ganglion and send information via long axons (either rostrally or caudally) to other ganglia. Through their patterns of outputs intersegmental interneurones are able to regulate the action of local circuits in different ganglia (Laurent and Burrows, 1989). In the metathoracic ganglion there are several populations of intersegmental interneurones with ascending or descending axons (Laurent and Burrows, 1988; Newland, 1990). Of the few interneurones studied in detail we know that they have similar exteroceptive receptive field properties to spiking local interneurones with specific arrays of mechanosensory hairs providing inputs to an interneurone (Newland and Burrows, 1997). We do not yet know, however, if they also receive and process chemosensory inputs from basiconic sensilla on the leg. In the sub-oesophageal ganglion there is considerable evidence to show that intersegmental interneurone receive mechanosensory inputs when basiconic sensilla on the palp dome are mechanically stimulated with plant material (Simpson, 1992). Few studies have analysed the properties of chemosensory inputs to intersegmental interneurone. Rogers (1998), despite characterizing several intersegmental neurones, failed to find any in the sub-oesophageal ganglion that responded differently to either various plant tissues or droplets of aqueous extractions of plant material applied to the palp domes. All neurones apparently responding to chemosensory stimulation in this study were local interneurones. Coordinated responses of the animal to stimulation of the tarsi and palps are essential in the control of feeding and other behaviours. Intersegmental interneurones are integral to such behaviours and must therefore play an important role in gustatory processing. It is clear that we must now analyse the responses of this important class of interneurones in order to understand their role in chemosensory behaviour.
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THE MOTOR OUTPUT OF LOCAL CIRCUITS
While the different classes of spiking interneurones are principally involved in sensory processing and coordination, non-spiking interneurones are primarily involved in motor control (Burrows, 1980). As their name implies, they communicate by means of graded electrical potentials, and by virtue of this exert a precise and continuous control over motor neurones. Unlike the two previous classes of spiking interneurones, the non-spiking interneurones receive fewer monosynaptic inputs from exteroceptive receptors (Laurent and Burrows, 1988). They do, however, receive monosynaptic inputs from spiking local interneurones, so that any mechano- or chemosensory drive to these interneurones will be specified by the action of presynaptic spiking local interneurones. An important feature of non-spiking interneurones that makes them key components in motor control is their divergent pattern of connections with different motor neurones. Each non-spiking interneurone can make synaptic connections with a number of motor neurones, and in turn each motor neurone receives convergent input from a number of non-spiking interneurones (Burrows, 1980). This organization means that the movements of a leg or mouthpart result from the orchestrated action of many interneurones and motor neurones. Based on many studies the design principles on which local circuits are organized can be described in detail (Fig. 10). This does not however mean that we necessarily know exactly what the output of a local circuit will be in
FIG. 10 Summary diagram of the basic organization of chemo- and mechanosensory processing pathways in local circuits. The coding of the neurone types filled in black is discussed in Section 5. Taken from Rogers and Newland (2002), ß 2002 by the Society for Neuroscience.
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response to a given sensory input. Neuronal signals are subject to a continuous modulation, so that the emergent outputs of these circuits will depend on the behavioural state of the animal, which may alter many factors including the membrane properties of the local circuit neurones, the synaptic weighting between individual components, inhibitory mechanisms and on the neuromodulators that are released. All of these factors are likely to be consequent on the nutritional status of an insect and thus subject to modification in the light of its previous feeding experience
5
Chemosensory coding in the metathoracic ganglion of the locust
In Sections 3 and 4 we summarized the anatomical and physiological evidence that local circuits in the metathoracic ganglion receive and process chemosensory inputs from basiconic sensilla on the hind leg in the locust. In this section we address the question of what kind of chemosensory coding is performed by these local circuits, how this information is used to cue behavioural responses, and what conclusions can be drawn about the central processing of contact-chemosensory inputs in insects. 5.1
BEHAVIOURAL RESPONSES TO CHEMOSENSORY STIMULATION
We have developed a simple behavioural assay to measure the type of behavioural response that can be elicited by chemosensilla on the hind leg of the locust (Rogers and Newland, 2000). Solutions of different chemicals are applied as droplets onto the hind tarsus and the response of the animal, if any, recorded. We found that the only responses that could be reliably elicited were rapid reflex withdrawal movements in which the leg was raised out of the solution and either held in the air or replaced on the ground in another location (Fig. 11), movements closely resembling those evoked by tactile stimulation. Water droplets elicited responses in approximately 10% of applications, but adding another chemical to the water droplet increased the likelihood of the locust responding in a concentration-dependent manner; the higher the chemical concentration in the droplet the greater the probability that the locust withdrew its leg, until responses occurred in 70–80% of test applications. This was true of all the chemicals tested, ranging from the secondary plant compound nicotine hydrogen tartrate, to sodium chloride, through to nutrient chemicals such as sucrose and the amino acid salt lysine glutamate (Fig. 12). All these chemicals are known to stimulate gustatory neurones in locusts (Blaney and Duckett, 1975; White and Chapman, 1990; Simpson et al., 1991). Although it might be anticipated that a secondary plant compound such as nicotine hydrogen tartrate would elicit an aversive withdrawal response (White and Chapman, 1990), it is at first appearance much more surprising that
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FIG. 11 Local leg avoidance reflex in response to chemical stimuli applied to a hind tarsus. An aqueous droplet of chemical solution was applied to a hind of a locust. Droplets may evoke a rapid (<1 s) removal of the leg from the stimulus, either lifted clear of the stimulus or replaced on the substrate in another position. The likelihood of a locust responding to the stimulus depends on both the identity of the chemical in the solution and its concentration.
nutrient chemicals should also do so. The responses of the locusts to these chemicals do differ in one key parameter, however, and this is the concentration at which they become effective at eliciting withdrawal responses. For example, a concentration of only 5 mM of nicotine is needed to produce a behavioural response in 50% of applications, whereas 50 mM NaCl and nearly 500 mM of sucrose or lysine glutamate are needed to have the same behavioural effectiveness (Fig. 12). Adding a sub-behavioural threshold concentration of sucrose (100 mM) to a 50 mM sodium chloride solution has the effect of diminishing the probability of the locust responding to the droplet when applied to the tarsus, suggesting that the probability of the animal responding to a given solution is governed by a non-linear combination of all the sensory signals received from the chemicals in the solution. This implies that inputs from sensory neurones responding to different chemicals cannot be simply additive (Rogers and Newland, 2000). This behavioural data provided us with a model of how local circuit neurones in the metathoracic ganglion use chemosensory inputs to generate and co-ordinate a response to the chemosensory information they receive. What is clear from our studies is that it appears likely that all chemical inputs to local circuits code for aversion at the level of the local response of the hind leg itself, and that different chemicals become aversive, or deterrent, at different concentrations. This then implies that both chemical identity and concentration are important determinants of the behavioural response. The likelihood of the locust responding increases with concentration for any chemical, but for any particular concentration, different chemicals are more or less likely to elicit withdrawal behaviour. This simple behavioural model may seem a long way from the contactchemosensory sampling processes that lead to the initiation of feeding, but we
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suggest it offers several advantages. First, the solutions used in the behavioural assay are exactly those used in the physiological analysis of local circuits. Thus, there is a more direct link between stimulus, physiology and behaviour in this model than in trying to determine the correlation between the gustatory effects of a chemical in a tip recording solution to its effect in a complex solid food. Changes in the apparent concentration of a chemical in a solid food or chemical mixture may alter its behavioural significance to the insect. Second, there was good evidence (Section 4) that chemosensory neurones on the leg provided a synaptic drive onto local neuronal circuits that we already knew a great deal about; therefore we could give a context to the flow of chemosensory information through the metathoracic ganglion. Third, the behavioural assay led to rapid and unambiguous behavioural responses of a kind that proved easy to detect in the local circuits of the metathoracic ganglion. Quantifying the effects of chemosensory inputs that exerted their effects more indirectly through, for example, the wide ranging modulation of connections throughout a neuronal network, such as is known to occur in snails and leeches, would be much more difficult. A disadvantage of the assay in its present form is that it only offers the possibility of looking at rejection responses. In undisturbed locusts searching for potential foods appetitive stimuli contacting the tarsi should lead to a quite separate set of behaviours involving lowering the head and further contactchemosensory sampling by the mouthparts. By definition this must involve the conveying chemosensory signals out of the thorax and up to the suboesophageal ganglion and/or brain by intersegmental interneurones. The identity and hence properties of this population of interneurones is as yet completely unknown.
5.2
RESPONSES OF SPIKING LOCAL INTERNEURONES TO DIFFERENT CHEMICAL SOLUTIONS
Spiking local interneurones, like basiconic sensilla on the leg and mouthparts, are bimodal, receiving both chemosensory and mechanosensory inputs (Newland, 1999). Given that the majority of taste stimuli will be in a form that will invariably also have a mechanosensory component, chemosensation in this system, and indeed in any other system that utilizes bimodal chemosensory/mechanosensory sensilla, can be viewed as a form of modified touch. Chemosensory inputs enhance, or perhaps indirectly diminish, a mechanosensory signal making a behavioural response more likely to happen (Simpson, 1992), truly contact-chemoreception. Rogers (1998) and Rogers and Simpson (1999) described a number of interneurones in the sub-oesophageal ganglion of the locust that were fundamentally mechanosensitive, responding strongly to any object touching the palp domes, but the responses were modified according to the chemical
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FIG. 12 Recordings from four different spiking local interneurones showing responses to repeated applications of solutions of A, Sodium Chloride, B, Sucrose, C, Nicotine Hydrogen Tartrate and D, Lysine Glutamate applied as droplets to a hind leg. Concentration ranges used are the same as those in the behavioural assay of leg withdrawal response. The duration of depolarization and number of action potentials evoked in the interneurones increased with concentration for all four chemicals. Based on Rogers and Newland (2002), ß 2002 by the Society for Neuroscience.
properties of the substance (plant tissue) in contact with the mouthparts. This suggests that the chemosensory processing of inputs from mouthpart sensilla may also be primarily bimodal with mechanosensory processing, and that the leg-based taste model we have developed may have a general applicability to
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contact-chemosensory processing by the mouthparts. This perhaps accords with early observations of food sampling behaviour in extremely food-deprived locusts in which it was thought that only mechanical and not chemical qualities were used to assess what would be bitten (Dadd, 1963). In extreme cases the chemosensory component becomes less important in the pathway and locusts bite anything that their mandibles can enclose. There is indirect evidence for similar bimodal processing in caterpillars. Touching the mouthparts elicited rhythmical chewing in Manduca sexta caterpillars in which the suboesophageal connectives had been cut, but food-plant sap triggered much longer-lasting chewing bouts (Rowell and Simpson, 1992). In flies there is also a report of some bimodally sensitive interneurones in the sub-oesophageal ganglion, responding to touch of the labellum, but with modified responses according to the chemical qualities of the contacting droplet (Mitchell and Itagaki, 1992). We have made numerous intracellular recordings of spiking local interneurones during stimulation of bimodal basiconic sensilla and tactile hairs on the leg with ascending concentration series of the same chemical solutions used in the behavioural assay (Rogers and Newland, 2002). Spiking local interneurones responded more strongly as chemical concentration increased for all of the chemicals tested. There was a significant correlation between chemical concentration and the duration and/or the number of action potentials produced in 80% of neurones tested (Figs. 12 and 13). The high likelihood of any local interneurone responding in a concentrationdependent manner to any one of the test chemicals applied to the leg strongly suggests that inputs from sensory neurones sensitive to all of these chemicals converge onto the same interneurones, as would be predicted from their anatomical organization (Newland et al., 2000; see Section 3). Local interneurones differed only in their somatosensory receptive field and in their overall sensitivity. Some interneurones were extremely phasic in response (e.g. Fig. 12B), some phaso-tonic (e.g. Fig. 12D), and others intermediate in character (Fig. 12A and C). We found no evidence that different interneurones were differentially sensitive to only one of the chemicals; individual interneurones responded in the same phasic or phaso-tonic manner to all four chemicals (Rogers and Newland, 2002). As with the behavioural responses, the concentration of chemical in a solution needed to evoke a response stronger than a purely mechanosensory stimulus differed considerably between chemicals and there was an extremely close correspondence between the likelihood of evoking a behavioural response and the relative duration of response in spiking local neurones (Fig. 13). This is particularly clear when neural responses are normalized to give the same mean duration of responses to water droplets. Given this transformation, the chemosensory response profiles match the behavioural data almost exactly.
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FIG. 13 The close correlation between the behavioural responses of locusts to the duration of depolarization evoked in spiking local interneurones to different chemicals applied to a hind leg. The X-axis shows chemical concentration (log10 scale), the left Y-axis the mean duration of depolarization in spiking local interneurones (black symbols) and the right Y-axis the proportion of free-moving locusts withdrawing their hind leg (white symbols) from a droplet of a chemical solution applied to the tarsus. The duration of neural response and the likelihood of behavioural withdrawal both increase similarly with chemical concentration, but different chemicals become effective stimuli at different concentration thresholds. Taken from Rogers and Newland (2002), ß 2002 by the Society for Neuroscience.
5.3
RESPONSES OF LEG MOTOR NEURONES
Recordings from the output stage of the metathoracic local circuits reveal that the pattern of neural sensitivity is maintained throughout the neuronal network (Rogers and Newland, 2002). The size of the synaptic inputs to leg motor neurones increases depending on chemical identity and concentration, just as in the behavioural model and the physiological responses of the spiking local interneurones. Figure 14 shows the responses of a flexor tibiae motor neurone (of which there are at least 9), which has an important role in generating the withdrawal behaviour, to an ascending concentration series of four different chemicals applied to the hind tarsus. Both the amplitude and the duration of the synaptic input increase with concentration. The antagonists of these motor neurones, the extensor tibiae motor neurones, receive an increasing inhibitory drive that mirrors the levels of excitation received by the flexor tibiae motor neurones.
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FIG. 14 The amplitude and duration of response of leg motor neurones increases with concentration according to chemical identity. Recording from a single flexor-tibiae motor neurone showing responses to all the chemical solutions used in the behavioural assay when applied to a hind leg. The increase in duration and amplitude of neural response parallels the probability of evoking a behavioural response to the different chemical solutions and resembles the responses seen in spiking local interneurones. Based on Rogers and Newland (2002), ß 2002 by the Society for Neuroscience.
Since spiking local interneurones in the midline group all contain the inhibitory transmitter GABA and therefore must have inhibitory outputs (see Section 4 above), they cannot directly mediate the excitatory responses of flexor tibiae motor neurones. At least one more, sign reversing, level of integration must be interposed, which could be generated through other populations of spiking local interneurones that form excitatory outputs with flexor motor neurones (Nagayama, 1989), or via non-spiking local interneurones that control sets of motor neurones (Burrows, 1980). The dose-dependent inhibitory responses of other motor neurones could, however, be the consequence of the direct inhibition by midline spiking local interneurones (Burrows and Siegler, 1982). The size of the synaptic input to motor neurones increases in a graded manner, but the behavioural response is all or nothing. As described in Section 4 above, the likelihood of a motor neurone depolarizing sufficiently to
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generate action potentials and hence move the leg is dependent not only on the input it receives ultimately from chemosensory neurones but also from other proprioceptive, intersegmental and neuromodulatory sources converging onto it (Burrows, 1996). A larger chemosensory input makes it more likely that a motor neurone will achieve spiking threshold, but does not guarantee it. The most remarkable feature of the local circuits in the metathoracic ganglion is that the basic organization of the behavioural output is almost fully apparent at a primary synaptic stage, that between chemosensory neurones and their target spiking local interneurones. There are several factors that will determine how contact-chemosensory information will be represented in these interneurones. The spiking frequency of chemosensory neurones and the strength of synapse they make onto local interneurones will be important components in determining the duration and amplitude of the response and hence the number of action potentials produced. The firing frequency of a chemosensory neurone will be affected by its sensitivity to particular chemicals, the concentration of those chemicals and the number of chemicals in a mixture, leading to interactions within and between chemosensory neurones. The size of post-synaptic potentials elicited by an individual action potential will be unitary in size for any particular chemosensory neurone, but may vary between sensory neurones synapsing onto the same interneurone, such that a strongly responding chemosensory neurone may only have a small effect on a particular interneurone compared to another sensory neurone producing fewer action potentials. Contact-chemosensory information is conveyed through the local circuits to motor neurones in a way that does not alter the relative size of the responses to the different chemical solutions, even though the sign of the signal may be reversed between antagonistic motor neurones. At a practical level this may offer some advantages when working on other insect species with less wellcharacterized central nervous systems than the locust. Motor neurones are generally more easily identified and characterized than local interneurones. Retrograde dye labelling similar to that used in establishing sensory projections can be used to identify motor neurones, which are frequently much larger and easier to record from than local interneurones. The key requirement in any analysis would be to establish behaviours that are clearly and rapidly influenced by contact-chemosensory input. Until recently, next to nothing was known about the neuronal networks that control feeding behaviour in insects and therefore it would be very difficult to establish how chemosensory signals affect these motor patterns. Small parts of the feeding process have been described. Mandibular motor neurones and mandibular premotor neurones, some of which are known to receive some form of sensory inputs from peripheral nerves have been described in the moth caterpillar Manduca sexta (Griss, 1990; Rohrbacher, 1994). Inhibitory thoracic inputs controlling chewing activity have also been described in caterpillars of
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Manduca sexta (Griss et al., 1991; Rowell and Simpson, 1992). In locusts, some elements of the central pattern generator controlling movements of the mouthparts have also been recently analysed (Rast, 2001; Rast and Bra¨unig, 2001) and the pattern of activity of the satellite nervous system during feeding and its possible role in controlling the salivary glands has been described (Bra¨unig, 1988, 1990; Schachtner and Bra¨unig, 1993, 1995). In general, the integration of gustatory signals onto neuronal networks controlling feeding is much better understood in other invertebrate models, where for example, the role of chemosensory inputs in promoting buccal mass and radular activity is well documented in molluscs (Delaney and Gelperin, 1990; Yoshida and Kobayashi, 1992; Elliott and Susswein, 2002), as have the chemosensory inputs onto the serotonergic Retzius cells of leeches that help initiate feeding motor patterns (Lent et al., 1989; Groome et al., 1995). 5.4
SIGNIFICANCE AND GENERAL APPLICABILITY OF THE LOCUST MODEL
Since the duration of response in spiking local interneurones in the metathoracic ganglion depends on both chemical identity and concentration, it is highly unlikely that these interneurones are able to code for chemical identity as such. Different combinations of chemical and concentration can produce exactly the same magnitude of response in any particular local interneurone (Fig. 13). It would therefore be impossible for any other population of neurones in the central nervous system to read out chemical identity from these responses based solely on this criterion. What useful information then could be extracted by the central nervous system using this sensory processing organization if not chemical identity? We believe that the evidence suggests that the local circuits are coding directly for a single chemical quality: aversiveness (Rogers and Newland, 2002). The relative size of neuronal response in spiking local interneurones and leg motor neurones closely parallels the behavioural likelihood of a locust removing its leg from a droplet depending on chemical identity and concentration (Fig. 13). It is irrelevant what the chemical is, only that there is too much of it present; this may be a very small amount for some secondary plant compounds, but considerably more in the case of nutrient chemicals. It is unsurprising that locusts should be highly sensitive to, and avoid toxic secondary plant compounds such as nicotine. It is perhaps less obvious at first why they should find higher concentrations of nutrient chemicals aversive. This behavioural and physiological response, however, shows some consistency with the model of the role of taste in dietary regulation by Simpson and Raubenheimer (1996) as discussed above in Section 2. To reiterate, in this model an animal does not necessarily have an open-ended appetitive desire for any, or all, nutrient chemicals, but instead these have to be consumed in the right relative proportions to achieve a balanced diet. Highly concentrated sources of single nutrients, therefore, are not only less desirable than more
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balanced blends of different nutrients but they may be actively rejected at the gustatory sampling stage prior to feeding. Recently a model with some similar features has been developed for taste processing in vertebrates, where interactions between the basic taste qualities provide overarching measures of two opposing qualities, nutritional suitability or toxicity (Scott and Giza, 2000; Scott and Verhagen, 2000). Our behavioural and neurophysiological data constitute a negative ‘taste model’ (Simpson and Raubenheimer, 1996), in which neural response and behavioural aversiveness increase with chemical concentration in potential foods, defining a region (the trough in Fig. 15A) in which items are not rejected and presumably suitable for further contact-chemosensory sampling. The metathoracic behavioural model is admittedly extreme, the locust rapidly lifts its leg from the applied droplet, but at lower concentrations the same information could make an insect less likely to stop at a potential food source and therefore never start sampling with its mouthparts, thereby rejecting the food source at an early stage. An important consequence of the contact-chemosensory organization described above is that individual chemicals cannot be considered intrinsically phagostimulatory and in this it contrasts strongly with the earlier models proposed by Dethier (1973) and Schoonhoven (1987). Any chemical is potentially phagodeterrent at the right concentration, and if not present in the correct balance with other chemicals. We do not wish to suggest that all contact-chemosensory processing in insects is solely governed by rejection, as there is of course much evidence for the phagostimulatory qualities of some chemicals, which actively promote feeding. The chemosensory corollary of palatability however may be governed by similar general principles as aversion. These principles are, first, that the central nervous system may be coding directly for a chemical quality or qualities rather than analysing for chemical identity and then making a decision on whether to feed. This quality will be dependent on a combination of chemical identity, concentration and blend (as in aversion coding) and relative palatability will be determined simply by the magnitude of an interneurone response that is a function of all of these factors. Second, chemosensory afferents will synapse directly onto and excite neuronal circuits that organize feeding-associated behaviours, which will also involve mechanosensory inputs. Finding these neuronal inputs may well prove more difficult than those coding for aversion. The hind leg system is reactive, the leg was in situ, the stimulus applied and the locust responded. Neuronal circuits that organize avoidance are more likely to remain receptive to sensory inputs at all times than neuronal circuits that initiate behaviours such as feeding, which are much more context, and nutritional state dependent. Furthermore, food sampling behaviours by the mouthparts are mostly proactive, they are actively brought to bear onto a surface to sample it. Although there have been no direct demonstrations of chemosensory inputs onto central pattern generators or
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FIG. 15 Interactions between local circuits organizing avoidance and acceptance behaviours. (A) Simple model of interaction between two local circuits. The aversion or rejection circuit is similar to that we have analysed in the metathoracic ganglion of the locust (in black). As yet there has been no experimental analysis of a local circuit organizing a feeding behaviour in insects (shown in grey). (B) Chemosensory quality coding in an interneurone receiving inputs from two different sensory neurones. The magnitude of the post-synaptic response increases according to chemical concentration of each of the chemicals. (C) Differences in dose–response curves of interneurones that are part of the rejection and acceptance local circuits. Because of inhibition from the rejection circuit feeding can only occur when there is a maximal difference between the two curves (arrow), which occurs at intermediate chemical concentrations. See text for full explanation.
other neuronal circuits associated with feeding behaviours in insects (only indirect evidence, e.g. Rowell and Simpson, 1992), such direct chemosensory inputs are well known in snails (Elphick et al., 1995; Kemenes et al., 2001), where synaptic inputs from sucrose-sensitive neurones promote activity in the neuronal circuits controlling feeding movements.
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The simplest overall contact-chemosensory organization in an insect is probably one that would combine a pathway that leads to an aversion or rejection behaviour coupled with a pathway that promotes a feeding behaviour. A simple scheme for combining these two pathways is shown in Fig. 15. In Fig. 15 two chemosensory neurones respond to chemicals X and Y respectively with simple sigmoid dose–response relationships. These chemicals are either nutrients or necessary to the correct host selection of the insect, but are not open-ended sign stimuli; ingested quantities of both have to be regulated. The sensory neurones make divergent synaptic connections onto interneurones R and A. Interneurone R is part of a circuit that organizes rejection or avoidance behaviours such as we have described above. This interneurone responds to chemicals X and Y in the manner shown in Fig. 15B, with the size of synaptic input increasing with concentration and varying with the blend of the two chemicals (cf. Fig. 13). Interneurone A is similar in many respects to interneurone R but synapses onto a neuronal circuit that organizes a feeding-associated (acceptance) behaviour and critically receives a stronger synaptic input from both sensory neurones. Interneurone A therefore has a greater sensitivity to the two chemicals, such that the dose–response curves seen in this interneurone start to rise at lower concentrations of the chemicals than in interneurone R, as shown in Fig. 15C. Low concentrations of chemical fail to elicit any response in either pathway. Increasing concentrations of the chemical more strongly stimulate the feeding pathway, but as concentration increases yet further also begin to stimulate the rejection pathway. At very high concentrations both rejection and feeding pathways are highly stimulated. It is a critical feature of the model, therefore, that the rejection pathway can strongly inhibit the acceptance pathway and prevent the transmission of information to the neuronal circuits that organize the feedingassociated behaviour. Thus at high chemical concentrations rejection is the dominant pathway. Feeding behaviours can occur when the inhibition from the interneurone R is minimal compared to the excitation in interneurone A and therefore the level of response in the two interneurones A and R are maximally different (arrow, Fig. 15C). The space defined by the differential responses of the two interneurones R and A in the two different pathways is likely to vary between chemicals. Sensory neurones sensitive to macronutrient and other major food indicator chemicals are likely to have much stronger synaptic inputs onto the acceptance than rejection pathways. For other chemicals, which may only be tolerable in small quantities the strengths of synaptic inputs onto the two pathways may be more similar. Whilst it is possible that all chemosensory neurones are completely cross-wired onto the two pathways, in some instances sensory neurones specifically sensitive to particularly aversive chemicals may only synapse onto the rejection pathway, and conversely for insects that do not take regulated meals, they may have sensory neurones sensitive to phagostimulants that only synapse onto an acceptance pathway
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(as proposed by Chapman, 2003). Apparently deterrent neurones may synapse onto both pathways. For example, some essential amino acids are known to stimulate deterrent neurones (e.g. Bernays and Chapman, 2001), which at first appears to make little sense, but if the sensory neurone responds at a moderate firing rate to the amino acid it may lead to stronger activity in an acceptance rather than rejection pathway and therefore contribute to the overall phagostimulatory input into the acceptance pathway. Another undoubtedly important component in any chemosensory processing pathway will be connections onto neuromodulatory neurones that will alter the characteristics of neuronal circuits controlling feeding or other behaviours. Again work on snails (Elphick et al., 1995) may indicate the kind of processes that could occur in the insect central nervous system, and may provide the neural substrate for the centrally excited state (Dethier, 1976) that promotes the continuation of feeding once it has started. Nitric oxide, for example, is known to activate the central pattern generator in the suboesophageal ganglion of locusts that controls the mandibles (Rast, 2001). Nutrient-specific feedbacks that modulate the responsiveness of sensory neurones to individual chemicals (as described in Section 2 above), perhaps in conjunction with centrally effective feedbacks will probably adjust the shape of the response profiles of individual interneurones and the relationship between neuronal circuits organizing different behaviours. These may serve to selectively raise or lower the thresholds of acceptance and rejection behaviours. These may be more important in neuronal circuits that control the final stages of food acceptance; chemosensory neurones on the legs of locusts, for example, do not show the same nutrient-specific feedbacks as those on the palps (Simpson, 1990).
6
Concluding remarks
There is still a degree of uncertainty about exactly what qualities of a stimulus contact-chemosensory neurones encode, whether its chemical identity is read out using across-fibre patterning, or a more abstract taste quality such as the vertebrate qualities of sweet, salty, sour and bitter, or more direct measures of phagostimulatory or aversive qualities. Modulation of the sensory response at the receptor level can have an important role in shaping the coding properties of sensory neurones, in regulating dietary intake and in the longterm setting of host plant preferences. Contact-chemosensory neurones mostly project to and arborize within their local ganglion or neuromere. There is no evidence at present for a dedicated gustatory processing region in the brain akin to the olfactory lobes. From sensory projection patterns it would appear that gustatory processing is carried out at multiple locations within the central nervous system. A minority of chemosensory neurones from mouthpart chemosensilla may send axons up
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into the brain where they arborize in a region adjacent to the antennal lobes. In the Orthoptera, chemosensory axons project to the same region of the ganglion as mechanosensory neurones originating from the same part of the leg. The available evidence, however, indicates that there is a modality-specific segregation of chemosensory and mechanosensory neurones in the central nervous system of flies. We have made pleasing progress in understanding the role of local neuronal circuits in chemosensory processing using the locust hind leg – metathoracic ganglion system of the locust as a model. The few studies on the central nervous processing of gustatory signals all indicate that interneurones receiving chemosensory inputs also receive mechanosensory inputs and are hence bimodal in function. Furthermore it is likely that individual interneurones receive inputs from multiple chemosensory neurones coding for different chemical sensitivities. In the metathoracic ganglion, chemo-mechanosensory interneurones appear to be coding directly for a chemical quality, that of aversiveness, which is a function of chemical identity, concentration and chemical blend. The sensitivities of individual chemosensory neurones, the interactions between them and the strength of synaptic connection they make onto target interneurones will all be key components in shaping the profile of this chemosensory quality. We speculate that other chemosensory qualities, associated with palatability may also be directly encoded in similar local circuits that organize feeding-associated behaviours. This organization contrasts strongly with that of the olfactory system in insects, where there is a high degree of anatomical chemotopy with different odorants activating a wide variety of combinations of interneurones (Hildebrand and Shepherd, 1997; Laurent, 1997; Hansson and Anton, 2000). It is of course possible that there are other contact-chemosensory processing pathways where more segregated information is used, or where responses of populations of interneurones could be used to distinguish one chemical from another. Given the generally distributed projections of chemosensory neurones, however we believe that direct contact-chemosensory connections onto local circuits, which do not need complex gustatory coding neuropils, will be common in the central nervous system, and do much to shape the feeding preferences of insects.
Acknowledgements This work has been principally funded by the Biotechnology and Biological Sciences Research Council (UK) whose support we gratefully acknowledge. We would like to thank Prof. Malcolm Burrows, and Drs Tom Matheson and Ibrahim Gaaboub for contributing to the work presented here. We are also grateful to numerous people in the Universities of Cambridge, Oxford and Southampton for many fruitful discussions on this work.
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References Abisgold, J. D. and Simpson, S. J. (1988). The effect of dietary protein levels and haemolymph composition on the sensitivity of the maxillary palp chemoreceptors of locusts. J. Exp. Biol. 135, 215–229. Akers, R. P. and Getz, W. M. (1993). Response of olfactory receptor neurones in honey bees to odorants and their binary mixtures. J. Comp. Physiol. A 173, 169–185. Altner, H. (1977). Insect sensillum specificity and structure: an approach to a new typology. In: Olfaction and Taste VI (eds. Magnen, J., and MacCleod, P.), pp. 295–303. London: Information Retrieval Ltd. Amakawa, T. (2001). Effects of age and blood sugar levels on the proboscis extension of the blow fly Phormia regina. J. Insect Physiol. 47, 195–203. Araneda, R. C., Kini, A. D. and Firestein, S. (2000). The molecular receptive range of an odor receptor. Nat. Neurosci. 3, 1248–1255. Ascoli-Christensen, A., Sutcliffe, J. F. and Albert, P. J. (1990). Effect of adenine nucleotides on labellar chemoreceptive cells of the stable fly Stomoxys calcitrans. J. Insect Physiol. 36, 339–344. Bacon, J. P. and Altman, J. S. (1977). A silver intensification method for cobalt-filled neurones in wholemount preparations. Brain Res. 138, 359–363. Bell, W. J. (1990). Searching behavior patterns in insects. Ann. Rev. Entomol. 35, 447–467. Bernays, E. A. and Chapman, R. F. (1972). The control of changes in peripheral sensilla associated with feeding in Locusta migratoria (L). J. Exp. Biol. 57, 755–763. Bernays, E. A. and Chapman, R. F. (1994). Host Plant Selection Behaviour of Phytophagous Insects. New York: Chapman and Hall. Bernays, E. A. and Chapman, R. F. (1998). Phenotypic plasticity in numbers of antennal chemoreceptors in a grasshopper: effects of food. J. Comp. Physiol. A 183, 69–76. Bernays, E. A. and Chapman, R. F. (2001). Taste cell responses in the polyphagous arctiid, Grammia geneura: towards a general pattern for caterpillars. J. Insect Physiol. 47, 1029–1043. Bernays, E. A. and Simpson, S. J. (1982). Control of food intake. Adv. Insect Physiol. 16, 59–102. Bernays, E. A. and Weiss, M. R. (1996). Induced food preferences in caterpillars: the need to identify mechanisms. Entomol. Exp. Appl. 78, 1–8. Bernays, E. A., Blaney, W. M. and Chapman, R. F. (1972). Changes in chemoreceptor sensilla on the maxillary palps of Locusta migratoria in relation to feeding. J. Exp. Biol. 57, 745–753. Bernays, E. A., Chapman, R. F. and Singer, M. S. (2000). Sensitivity to chemically diverse phagostimulants in a single gustatory neurone of a polyphagous caterpillar. J. Comp. Physiol. 186, 13–19. Blaney, W. M. (1975). Behavioural and electrophysiological studies of taste discrimination by the maxillary palps of larvae of Locusta migratoria (L). J. Exp. Biol. 62, 555–569. Blaney, W. M. (1977). The ultrastructure of an olfactory sensillum on the maxillary palps of Locusta migratoria. Cell Tissue Res. 184, 397–409. Blaney, W. M. (1981). Chemoreception and food selection in locusts. Trends Neurosci. 3, 35–39. Blaney, W. M. and Chapman, R. F. (1969). The anatomy and histology of the maxillary palp of Schistocerca gregaria (Orthoptera, Acrididae). J. Zool. 157, 509–535. Blaney, W. M. and Chapman, R. F. (1970). The functions of the maxillary palps of acrididae (orthoptera). Entomol. Exp. Appl. 13, 363–376.
THE NEUROBIOLOGY OF TASTE IN INSECTS
195
Blaney, W. M. and Duckett, A. M. (1975). The significance of palpation of the maxillary palps of Locusta migratoria (L): an electrophysiological and behavioural study. J. Exp. Biol. 63, 701–712. Blaney, W. M. and Simmonds, M. S. J. (1985). Food selection by locusts: the role of learning in rejection behaviour. Entomol. Exp. Appl. 39, 273–278. Blaney, W. M. and Simmonds, M. S. J. (1990a). The chemoreceptors. In: Biology of Grasshoppers (eds. Chapman, R. F., and Joern, A.), pp. 1–37. New York: John Wiley and Sons. Blaney, W. M. and Simmonds, M. S. J. (1990b). A behavioural and electrophysiological study of the role of tarsal chemoreceptors in feeding by adults of Spodoptera, Heliothis virescens and Helicoverpa armigera. J. Insect Physiol. 36, 743–756. Blaney, W. M. and Winstanley, C. (1980). Chemosensory mechanisms of locusts in relation to feeding: the role of some secondary plant compounds. In: Insect Neurobiology and Pesticide Action (Neurotox 79), pp. 383–389. London: Chem Industry. Blaney, W. M. and Winstanley, C. (1982). Food selection behaviour in Locusta migratoria. In: Proceedings of the 5th International Symposium on Insect-Plant Relations (eds. Visser, J. H., and Minks, A. K.), pp. 365–366. Wageningen: Pudoc. Blaney, W. M., Winstanley, C. and Simmonds, M. S. J. (1985). Food selection by locusts: an analysis of rejection behaviour. Entomol. Exp. Appl. 38, 35–40. Blom, F. (1978). Sensory activity and food intake: a study of input-output relationships in two phytophagous insects. Neth. J. Zool. 28, 277–340. Boeckh, J. and Ernst, K.-D. (1987). Contribution of single unit analysis in insects to an understanding of olfactory function. J. Comp. Physiol. A 161, 549–565. Bogner, F., Boppre, M., Ernst, K. D. and Boeckh, J. (1986). Carbon dioxide sensitive receptors on labial palps of Rhodogastria moths (Lepidoptera: Arctiidae): physiology, fine structure and central projection. J. Comp. Physiol. A 158, 741–750. Bowen, M. F., Davis, E. and Haggart, D. A. (1988). A behavioural and sensory analysis of host-seeking behaviour in diapausing mosquito Culex pipiens. J. Insect Physiol. 34, 805–813. Brandes, C., Frisch, B. and Menzel, R. (1988). Time-course of memory formation differs in honeybee lines selected for good and bad learning. Animal Behav. 36, 981–985. Bra¨unig, P. (1988). Neurosecretory cells of the locust suboesophageal ganglion. Symp. Biol. Hungarica 36, 161–171. Bra¨unig, P. (1990). The mandibular ganglion – a new peripheral ganglion of the locust. J. Exp. Biol. 148, 313–324. Bra¨unig, P., Hustert, R. and Pflu¨ger, H.-J. (1981). Distribution and specific central projections of mechanoreceptors in the thorax and proximal leg joints of locusts.1. Morphology, location and innervation of internal proprioceptors of prothorax and metathorax and their central projection. Cell Tissue Res. 216, 57–77. Brown, A. G., Rose, P. K. and Snow, P. J. (1977). The morphology of hair follicle afferent fibre collaterals in the spinal cord of the cat. J. Physiol. 272, 779–797. Brown, A. G., Fyffe, R. E. W., Noble, R., Rose, P. K. and Snow, P. J. (1980). The density, distribution and topographical organization of spinocervical tract neurones in the cat. J. Physiol. 300, 409–428. Burrows, M. (1980). The control of sets of motor neurones by local interneurones in the locust. J. Physiol. Lond. 298, 213–233. Burrows, M. (1987). Parallel processing of proprioceptive signals by spiking local interneurones and motor neurones in the locust. J. Neurosci. 7, 1064–1080.
196
S. M. ROGERS AND P. L. NEWLAND
Burrows, M. (1992). Reliability and effectiveness of transmission from exteroceptive sensory neurones to spiking local interneurones in the locust. J. Neurosci. 12, 1477–1489. Burrows, M. (1996). The Neurobiology of an Insect Brain. Oxford: Oxford University Press. Burrows, M. and Newland, P. L. (1993). Correlation between the receptive fields of locust interneurones their dendritic morphology, and the central projections of mechanosensory neurones. J. Comp. Neurol. 329, 412–426. Burrows, M. and Newland, P. L. (1994). Convergence of mechanosensory afferents from different classes of exteroceptors onto spiking local interneurones in the locust. J. Neurosci. 14, 3341–3350. Burrows, M. and Siegler, M. V. S. (1982). Spiking local interneurones mediate local reflexes. Science 217, 650–652. Burrows, M. and Siegler, M. V. S. (1984). The diversity and receptive fields of spiking local interneurones in the locust metathoracic ganglion. J. Comp. Physiol. A 224, 483–508. Chapman, R. F. (1982). Chemoreception: the significance of sensillum numbers. Adv. Insect Physiol. 16, 247–356. Chapman, R. F. (1988). Sensory aspects of host-plant recognition by Acridoidea: questions associated with the multiplicity of receptors and variability of response. J. Insect Physiol. 34, 167–174. Chapman, R. F. (1995). Chemosensory regulation of feeding. In: Regulatory Mechanisms in Insect Feeding (eds. Chapman, R. F., and De Boer, G.), pp. 101–136. New York: Chapman & Hall. Chapman, R. F. (2003). Contact chemoreception in feeding by phytophagous insects. Annu. Rev. Entomol. 48, 455–484. Chapman, R. F. and Fraser, J. (1989). The chemosensory system of the monophagous grasshopper, Bootettix argentatus Bruner (Orthoptera: Acrididae). Int. J. Insect Morph. Embryol. 18, 111–118. Chapman, R. F. and Lee, J. C. (1991). Environmental effects on numbers of peripheral chemoreceptors on the antennae of a grasshopper. Chem. Senses 16, 607–616. Chapman, R. F. and Thomas, J. G. (1978). The numbers and distribution of sensilla on the mouthparts of Acridoidea. Acrida 7, 115–148. Chapman, R. F., Ascoli-Christensen, A. and White, P. R. (1991). Sensory coding for feeding deterrence in the grasshopper Schistocerca americana. J. Exp. Biol. 158, 241–259. Christensen, T. A. and Hildebrand, J. G. (2002). Pheromonal and host-odor processing in the insect antennal lobe: how different? Curr. Opin. Neurobiol. 12, 393–399. Clements, A. N. (1992). The Biology of Mosquitoes, Vol. 1. London: Chapman and Hall. Clyne, P. J., Warr, C. G. and Carlson, J. R. (2000). Candidate taste receptors in Drosophila. Science 287, 1830–1834. Consoulas, C., Kent, K. S. and Levine, R. B. (1996). Remodeling of the peripheral processes and presynaptic terminals of leg motoneurones during metamorphosis of the hawk moth Manduca. J. Comp. Neurol. 372, 415–434. Dadd, R. H. (1963). Feeding behaviour and nutrition in grasshoppers and locusts. Adv. Insect Physiol. 1, 47–111. de Boer, G. (1993). Plasticity in food preference and diet-induced differential weighting of chemosensory information in larval Manduca Sexta. J. Insect Physiol. 39, 17–24. de Bruyne, M., Clyne, P. J. and Carlson, J. R. (1999). Odor coding in a model olfactory organ: The Drosophila maxillary palp. J. Neurosci. 19, 4520–4532. de Jong, R. and Kaiser, L. (1991). Odor learning by Leptopilina boulardi, a specialist parsitoid (Hymenoptera: Eucoilidae). J. Insect. Behav. 4, 743–750.
THE NEUROBIOLOGY OF TASTE IN INSECTS
197
de Jong, R. and Pham-Delegue, M. H. (1991). Electroantennogram response related to olfactory conditioning in the honey bee (Apis mellifera ligustica). J. Insect Physiol. 37, 319–324. Delaney, K. and Gelperin, A. (1990). Cerebral interneurones controlling fictive feeding in Limax maximas. III. Integration of sensory inputs. J. Comp. Physiol. A. 166, 327–343. del Campo, M. L., Miles, C. I., Schroeder, F. C., Mueller, C., Booker, R. and Renwick, J. A. (2001). Host recognition by the tobacco hornworm is mediated by a host plant compound. Nature 411, 186–189. den Otter, C. J., Tchicaya, T. and Schutte, A. M. (1991). Effects of age, sex and hunger on the antennal olfactory sensitivity of tsetse flies. Physiol. Entomol. 16, 173–182. Derridj, S., Wu, B. R., Stammitty, L., Garrec, J. P. and Derrien, A. (1996). Chemicals on the leaf surface, information about the plant available to insects. Entomol. Exp. Appl. 80, 197–201. Dethier, V. G. (1973). Electrophysiological studies of gustation in lepidopterous larvae. II. Taste spectra in relation to food-plant discrimination. J. Comp. Physiol. 82, 103–134. Dethier, V. G. (1976). The Hungry Fly. Cambridge, Mass: Harvard University Press. Dethier, V. G. and Bowdan, E. (1989). The effect of alkaloids on sugar receptors and feeding behaviour of the blowfly. Physiol. Entomol. 14, 127–136. Dethier, V. G. and Crnjar, R. M. (1982). Candidate codes in the gustatory system of caterpillars. J. Gen. Physiol. 79, 549–569. Devitt, B. D. and Smith, J. J. B. (1982). Morphology and fine-structure of mouthpart sensilla in the dark-sided cutworm Euxoa-messoria (harris) (Lepidoptera, Noctuidae). Int. J. Insect Morphol. 11, 255–270. Distler, P. G. and Boeckh, J. (1997). Central projections of the maxillary and antennal nerves in the mosquito Aedes aegypti. J. Exp. Biol. 200, 1873–1879. Du, Y. J., van Loon, J. J. A. and Renwick, J. A. A. (1995). Contact chemoreception of oviposition-stimulating glucosinolates and an oviposition deterrent cardenolide in two sub-species of Pieris napi. Physiol. Entomol. 20, 164–174. Dunipace, L., Meister, S., McNealy, C. and Amrein, H. (2001). Spatially restricted expression of candidate taste receptors in the Drosophila gustatory system. Current Biology 11, 822–835. Edgecomb, R. S. and Murdock, L. L. (1992). Central projections of axons from taste hairs on the labellum and tarsi of the blowfly, Phormia regina Meigen. J. Comp. Neurol. 315, 431–444. Elliott, C. J. H. and Susswein, A. J. (2002). Comparative neuroethology of feeding control in molluscs. J. Exp. Biol. 205, 877–896. Elphick, M. R., Kemenes, G., Staras, K. and O’shea, M. (1995). Behavioral role for nitric-oxide in chemosensory activation of feeding in a mollusk. J. Neurosci. 15, 7653–7664. Ernst, K. D., Boeckh, J. and Boeckh, V. (1977). A neuroanatomical study on the organization of the Central antennal pathways in insects. Cell Tissue Res. 176, 285–308. Feeny, P. (1991). Chemical constraints on the evolution of swallowtail butterflies. In: Plant-Animal Interactions: Evolutionary Ecology in Tropical and Temperate Regions (eds. Price, P. W., Lewinsohn, T. M., Fernandes, G. W., and Benson, W. W.), pp. 15–34. New York: John Wiley. Feeny, P., Sta¨dler, E., Ahman, I. and Carter, M. (1989). Effects of plant odor on oviposition by the black swallowtail butterfly, Papillo polyxenes (Lepidoptera: Papillionidae). J. Insect Behav. 2, 803–828.
198
S. M. ROGERS AND P. L. NEWLAND
Fiala, V., Glad, C., Martin, M., Jolivet, E. and Derridj, S. (1990). Occurrence of soluble carbohydrates on the phylloplane of maize (Zea mays L): variations in relation to leaf heterogeneity and position on the plant. New Phytologist. 115, 609–615. Fox, A. N., Pitts, R. J. and Zwiebel, L. J. (2002). A cluster of candidate odorant receptors from the malaria vector mosquito, Anopheles gambiae. Chem. Senses 27, 453–459. Getz, W. M. and Akers, R. P. (1994). Honeybee olfactory sensilla behave as integrated processing units. Behav. Neural Biol. 61, 191–195. Getz, W. M. and Akers, R. P. (1997). Response of American cockroach (Periplaneta americana) olfactory receptors to selected alcohol odorants and their binary combinations. J. Comp. Physiol. A 180, 701–709. Glendinning, J. I., Ensslen, S., Eisenberg, M. E. and Weiskopf, P. (1999). Diet-induced plasticity in the taste system of an insect: localization to a single transduction pathway in an identified taste cell. J. Exp. Biol. 202, 2091–2102. Glendinning, J. I., Brown, H., Capoor, M., Davis, A., Gbedemah, A. and Long, E. (2001). A peripheral mechanism for behavioral adaptation to specific ‘bitter’ taste stimuli in an insect. J. Neurosci. 21, 3688–3696. Glendinning, J. I., Davis, A. and Ramaswamy, S. (2002). Contribution of different taste cells and signalling pathways to the discrimination of ‘bitter’ taste stimuli by an insect. J. Neurosci. 22, 7281–7287. Gonka, M. D., Laurie, T. J. and Prete, F. R. (1999). Responses of movement-sensitive visual interneurones to prey-like stimuli in the praying mantis Sphodromantis lineola (Burmeister). Brain Behav. Evolut. 54, 243–262. Greenwood, M. and Chapman, R. F. (1984). Differences in numbers of sensilla on the antennae of solitarious and gregarious Locusta migratoria L. (Orthoptera: Gr Acrididae). Int. J. Insect Morph. Embryol. 13, 295–301. Griss, C. (1990). Mandibular motor neurones of the caterpillar of the hawk moth Manduca Sexta. J. Comp. Neurol. 296, 393–402. Griss, C., Simpson, S. J., Rohrbacher, J. and Rowell, C. H. F. (1991). Localisation in the central nervous system of larval Manduca sexta (Lepidoptera: Sphingidae) of areas responsible for aspects of feeding behaviour. J. Insect Physiol. 37, 477–482. Groome, J. R., Vaughan, D. K. and Lent, C. M. (1995). Ingestive sensory inputs excite serotonin effector neurones and promote serotonin depletion from the leech central nervous system and periphery. J. Exp. Biol. 198, 1233–1242. Hammer, M. (1993). An identified neurone mediates the unconditioned stimulus in associative olfactory learning in honeybees. Nature 366, 59–63. Hansson, B. S. and Anton, S. (2000). Function and morphology of the antennal lobe: New developments. Annu. Rev. Entomol. 45, 203–231. Hildebrand, J. G. (1995). Analysis of chemical signals by nervous systems. Proc. Natl. Acad. Sci. USA 92, 67–74. Hildebrand, J. G. (1996). Olfactory control of behaviour in moths: central processing of odor information and the functional significance of olfactory glomeruli. J. Comp. Physiol. A 178, 5–19. Hildebrand, J. G. and Shepherd, G. M. (1997). Mechanisms for olfactory discrimination: converging evidence for common principles across phyla. Ann. Rev. Neurosci. 20, 595–631. Hodgson, E. S., Lettvin, J. Y. and Roeder, K. D. (1955). Physiology of a primary chemoreceptor unit. Science 122, 417–418. Homberg, U., Christensen, T. A. and Hildebrand, J. G. (1989). Structure and function of the deutocerebrum in insects. Annu. Rev. Neurobiol. 34, 477–501. Jacobs, K. M., Mark, G. P. and Scott, T. R. (1988). Taste responses in the nucleus tractus solitarious of sodium-deprived rats. J. Physiol. (Cam.) 406, 393–410.
THE NEUROBIOLOGY OF TASTE IN INSECTS
199
Johnson, S. E. and Murphey, R. K. (1985). The afferent projection of mesothoracic bristle hairs in the cricket, Acheta domesticus. J. Comp. Physiol. A 156, 369–379. Kemenes, G., Staras, K. and Benjamin, P. R. (2001). Multiple types of control by identified interneurones in a sensory-activated rhythmic motor pattern. J. Neurosci. 21, 2903–2911. Kendall, M. D. (1970). The anatomy of the tarsi of Schistocerca gregaria Forska˚l. Z. Zellforsch. 109, 112–137. Kent, K. S. and Hildebrand, J. G. (1987). Cephalic sensory pathways in the nervous system of larval Manduca Sexta (Lepidoptera: Spingidae). Phil. Trans. R. Soc. Lond. B 315, 1–36. Kent, K. S. and Levine, R. B. (1988). Neural control of leg movements in a metamorphic insect: sensory and motor elements of the larval thoracic legs in Manduca Sexta. J. Comp. Neurol. 271, 559–576. Klein, U. (1981). Sensilla of the cricket palp: fine structure and spatial organization. Cell Tissue Res. 219, 229–252. Kral, K. (1998). Side-to-side head movements to obtain motion depth cues: A short review of research on the praying mantis. Behav. Process 43, 71–77. Kral, K. and Devetak, D. (1999). The visual orientation strategies of Mantis religiosa and Empusa fasciata reflect differences in the structure of their visual surroundings. J. Insect Behav. 12, 737–752. Kupfermann, I. (1994). Neural control of feeding. Curr. Opin. Neurobiol. 4, 869–876. Laurent, G. (1996). Dynamical representation of odors by oscillating and evolving neural assemblies. Trends Neurosci. 19, 489–496. Laurent, G. (1997). Olfactory processing: maps, time and codes. Curr. Opin. Neurobiol. 7, 547–553. Laurent, G. and Burrows, M. (1988). Direct excitation of nonspiking local interneurones by exteroceptors underlies tactile reflexes in the locust. J. Comp. Physiol. A 162, 563–572. Laurent, G. and Burrows, M. (1989). Intersegmental interneurones can control the gain of reflexes in adjacent segments of the locust by their action on nonspiking local interneurones. J. Neurosci. 9, 3030–3039. Laurent, G., Stopfer, M., Friedrich, R. W., Rabinovich, M. I., Volkovskii, A. and Abarbanel, H. D. I. (2001). Odor encoding as an active, dynamical process: Experiments, computation, and theory. Annu. Rev. Neurosci. 24, 263–297. Lee, J. K. and Altner, H. (1986). Primary sensory projections of the labial palp-pit organ of Pieris rapae L (Lepidoptera: Pieridae). Int. J. Insect. Morphol. Embryol. 15, 439–448. Lent, C. M., Dickinson, M. H. and Cameron, G. M. (1989). Serotonin and leech feeding behaviour: obligatory neuromodulation. Am. Zool. 29, 1241–1254. Levine, R. B., Pak, C. and Linn, D. (1985). The structure, function and metamorphic reorganization of insect sensory systems. J. Comp. Physiol. A 157, 1–13. Lewis, C. T. (1970). Structure and function in some external receptors. Symp. R. Entomol. Soc. Lond. 5, 59–76. Ma, W. C. (1972). Dynamics of feeding responses in Pieris brassicae Linn. as a function of chemosensory input – a behavioural, ultrastructural and electrophysiological study. Meded. Landbouwhogesch. Wageningen. 72, 11–29. Masson, C. and Mustaparta, H. (1990). Chemical information processing in the olfactory system of insects. Physiol. Rev. 70, 199–245. Mitchell, B. K. (1987). Interactions of alkaloids with galeal chemosensory cells of the Colorado potato beetle. J. Chem. Ecol. 13, 2009–2022.
200
S. M. ROGERS AND P. L. NEWLAND
Mitchell, B. K. and Itagaki, H. (1992). Interneurones of the subesophageal ganglion of Sarcophaga bullata responding to gustatory and mechanosensory stimuli. J. Comp. Physiol. A 171, 213–230. Mitchell, B. K. and McCashin, B. G. (1994). Tasting green leaf volatiles by larvae and adults of Colorado potato beetles, Leptinotarsa decemlineata. J. Chem. Ecol. 20, 753–769. Mordue (Luntz), A. J. (1979). The role of the maxillary and labial palps in the feeding behaviour of Schistocera gregaria. Entomol. Exp. Appl. 25, 279–288. Mu¨cke, A. and Lakes-Harlan, R. (1995). Central projections of sensory cells of the midleg of the locust, Schistocerca gregaria. Cell Tissue Res. 280, 391–400. Murphey, R. K. (1981). The structure and development of a somatotopic map in crickets: the cercal afferent projection. Dev. Biol. 88, 236–246. Murphey, R. K., Jacklet, A. and Schuster, L. (1980). A topographic map of sensory cell terminal arborization in the cricket CNS: Correlations with birthday and position in a sensory array. J. Comp. Physiol. A 191, 53–64. Murphey, R. K., Possidente, D., Pollack, G. and Merritt, D. (1989a). Modality-specific axonal projections in the CNS of the flies Phormia and Drosophila. J. Comp. Neurol. 290, 185–200. Murphey, R. K., Possidente, D. R., Vandervorst, P. and Ghysen, A. (1989b). Compartments and the topography of leg afferent projections in Drosophila. J. Neurosci. 9, 3209–3217. Nagayama, T. (1989). Morphology of a new population of local interneurones in the locust metathoracic ganglion. J. Comp. Neurol. 283, 189–211. Naresh Singh, R. and Nayak, S. V. (1985). Fine structure and primary projections of sensilla on the maxillary palp of Drosophila melanogaster Meigen (Diptera: Drosophilidae). J. Insect Morphol. Embryol. 14, 291–306. Newland, P. L. (1990). The morphology of a population of ascending interneurones in the metathoracic ganglion of the locust. J. Comp. Neurol. 299, 242–260. Newland, P. L. (1991). Morphology and somatotopic organization of the central projections of afferents from tactile hairs on the hind leg of the locust. J. Comp. Neurol. 311, 1–16. Newland, P. L. (1998). Avoidance reflexes mediated by contact-chemoreceptors on the legs of locusts. J. Comp. Physiol. A 183, 313–324. Newland, P. L. (1999). Processing of gustatory information by spiking local interneurones in the locust. J. Neurophysiol. 82, 3149–3159. Newland, P. L. and Burrows, M. (1994). Processing of mechanosensory information from gustatory receptors on a hind leg of the locust. J. Comp. Physiol. A 174, 399–410. Newland, P. L. and Burrows, M. (1997). Processing of tactile information in neuronal networks controlling leg movements of the locust. J. Insect Physiol. 43, 107–128. Newland, P. L. and Emptage, N. J. (1996). The central connection and actions during walking of tibial campaniform sensilla in the locust. J. Comp. Physiol. A 178, 749–763. Newland, P. L., Rogers, S. M., Gaaboub, I. and Matheson, T. (2000). Parallel somatotopic maps of gustatory and mechanosensory neurones in the central nervous system of an insect. J. Comp. Neurol. 425, 82–96. Nottingham, S. F., Hardie, J., Dawson, G. W., Hick, A. J., Pickett, J. A., Wadhams, L. J. and Woodcock, C. M. (1991). Behavioural and electrophysiological responses of aphids to host and non-host plant volatiles. J. Chem. Ecol. 17, 1231–1242. Oldfield, B. P. (1982). Tonotopic organization of auditory receptors in Tettigoniidae (Orthoptera: Ensifera). J. Comp. Physiol. A 147, 461–469.
THE NEUROBIOLOGY OF TASTE IN INSECTS
201
Peterson, B. A. and Weeks, J. C. (1988). Somatotopic mapping of sensory neurones innervating mechanosensory hairs on the larval prolegs of Manduca Sexta. J. Comp. Neurol. 275, 128–144. Pfaffmann, C. (1941). Gustatory afferent impulses. J. Cell. Comp. Physiol. 17, 243–258. Pflu¨ger, H. J. (1980). The function of hair sensilla on the locust’s leg: the role of tibial hairs. J. Exp. Biol. 87, 163–175. Pflu¨ger, H. J., Bra¨unig, P. and Hustert, R. (1981). Distribution and specific central projections of mechanoreceptors in the thorax and proximal leg joints of locusts. II. The external mechanoreceptors: hair plates and tactile hairs. Cell Tissue Res. 216, 79–96. Pflu¨ger, H. J., Bra¨unig, P. and Hustert, R. (1988). The organization of mechanosensory neuropiles in locust thoracic ganglia. Phil. Trans. R. Soc. Lond. 321, 1–26. Pitman, R. M., Tweedle, C. D. and Cohen, M. J. (1972). Branching of central neurones: intracellular cobalt injection for light and electron microscopy. Science 176, 412–414. Prakash, S., Mendki, M. J., Rao, K. M., Singh, K. and Singh, R. N. (1995). Sensilla on the maxillary and labial palps of the cockroach Supella longipalpa Fabricus (Dictyoptera: Blattellidae). Int. J. Insect Morphol. Embryol. 24, 13–34. Rast, G. (2001). Nitric oxide induces centrally generated motor patterns in the locust suboesophageal ganglion. J. Exp. Biol. 204, 3789–3801. Rast, G. F. and Bra¨unig, P. (2001). Insect mouthpart motor patterns: Central circuits modified for highly derived appendages? Neurosci. 108, 167–176. Raubenheimer, D. and Simpson, S. J. (1993). The geometry of compensatory feeding in the locust. Anim. Behav. 45, 953–964. Robertson, H. M. (2001). Taste: independent origins of chemoreception coding systems. Current Biology 11, R560–R562. Roessingh, P., Sta¨dler, E., Scho¨ni, R. and Feeny, P. (1991). Tarsal contactchemoreceptors of the black swallowtail butterfly Papillo polyxenes: responses to phytochemicals from host- and non-host plants. Physiol. Entomol. 16, 485–495. Rogers, S. M. (1998). Chemosensory development and integration in Locusta migratoria. D.Phil. thesis, University of Oxford, UK. Rogers, S. M. and Newland, P. L. (2000). Local movements evoked by chemical stimulation of the hind leg in the locust Schistocerca gregaria. J. Exp Biol. 203, 423–433. Rogers, S. M. and Newland, P. L. (2002). Gustatory processing in thoracic local circuits of locusts. J. Neurosci. 22, 8324–8333. Rogers, S. M. and Simpson, S. J. (1997). Experience-dependent changes in the number of chemosensory sensilla on the mouthparts and antennae of Locusta migratoria. J. Exp. Biol. 200, 2313–2321. Rogers, S. M. and Simpson, S. J. (1999). Chemo-discriminatory neurones in the suboesophageal ganglion of Locusta migratoria. Entomol. Exp. Appl. 91, 19–28. Rohrbacher, J. (1994). Mandibular premotor interneurones of larval Manduca Sexta. J. Comp. Physiol. A. 175, 619–628. Rolls, E. T. (1989). Information processing in the taste system of primates. J. Exp. Biol. 146, 141–164. Ro¨mer, H. (1983). Tonotopic organization of the auditory neuropile in the bushcricket Tettigonia viridissima. Nature 306, 60–62. Ro¨mer, H., Marquart, V. and Hardt, M. (1988). Organization of a sensory neuropile in the auditory pathway of two groups of Orthoptera. J. Comp. Neurol. 275, 201–215. Rowell, C. H. F. and Simpson, S. J. (1992). A peripheral input of thoracic origin inhibits chewing movements in the larvae of Manduca Sexta. J. Insect. Physiol. 38, 475–483.
202
S. M. ROGERS AND P. L. NEWLAND
Schachtner, J. and Bra¨unig, P. (1993). The activity pattern of identified neurosecretory cells during feeding behaviour in the locust. J. Exp. Biol. 185, 287–303. Schachtner, J. and Bra¨unig, P. (1995). Activity pattern of suboesophageal ganglion cells innervating the salivary glands of the locust Locusta migratoria. J. Comp Physiol. A 176, 491–501. Schoonhoven, L. M. (1973). Plant recognition by lepidopterous larvae. Symp. R. Entomol. Soc. Lond. 6, 87–99. Schoonhoven, L. M. (1987). What makes a caterpillar eat? The sensory code underlying feeding behaviour. In: Perspectives in Chemoreception and Behaviour (eds. Chapman, R. F., Bernays, E. A., and Stoffolano, J. G.), pp. 69–97. New York: Springer-Verlag. Schoonhoven, L. M. (1996). After the Verschaffelt-Dethier era: the insect-plant field comes of age. Entomol. Exp. Appl. 80, 1–5. Schoonhoven, L. M. and Blom, F. (1988). Chemoreception and feeding behaviour in a caterpillar: towards a model of brain functioning in insects. Entomol. Exp. Appl. 49, 123–129. Schoonhoven, L. M. and van Loon, J. J. A. (2002). An inventory of taste in caterpillars: Each species its own key. Acta Zool. Acad. Sci. H 48, 215–263. Schoonhoven, L. M., Blaney, W. M. and Simmonds, M. S. J. (1992). Sensory coding of feeding deterrents in phytophagous insects. In: Insect-Plant Interactions, Vol. 4. (ed. Bernays, E. A.), pp. 59–79. Boca Raton, Florida: CRC press. Scott, T. R. and Giza, B. K. (2000). Issues of gustatory neural coding: Where they stand today. Physiol. Behav. 69, 65–76. Scott, T. R. and Verhagen, J. V. (2000). Taste as a factor in the management of nutrition. Nutrition 16, 874–885. Selzer, R. (1984). On the specificities of antennal olfactory cells of Periplaneta americana. Chem. Senses 8, 375–395. Shanbhag, S. R. and Singh, R. N. (1992). Functional implications of the projections of neurones from individual labellar sensillum of Drosophila melanogaster as revealed by neuronal-marker horseradish peroxidase. Cell Tissue Res. 267, 273–282. Siegler, M. V. S. and Burrows, M. (1983). Spiking local interneurones as primary integrators of mechanosensory information in locust. J. Neurophysiol. 50, 1281–1295. Siegler, M. V. S. and Burrows, M. (1984). The morphology of two groups of spiking local interneurones in the metathoracic ganglion of the locust. J. Neurosci. 6, 507–513. Simmonds, M. S. J. and Blaney, W. M. (1990). Gustatory codes in lepidopterous larvae. Symposia Biologica Hungarica 39, 17–27. Simmonds, M. S. J., Schoonhoven, L. M. and Blaney, W. M. (1991). Daily changes in the responsiveness of taste receptors correlate with feeding behaviour in larvae of Spodoptera littoralis. Entomol. Exp. Appl. 61, 73–82. Simmonds, M. S. J., Simpson, S. J. and Blaney, W. M. (1992). Dietary selection behaviour in Spodoptera littoralis: The effects of conditioning diet and conditioning period on neural responsiveness and selection behaviour. J. Exp. Biol. 162, 73–90. Simpson, C. L. (1990). Dietary compensation by Locusta migratoria: aspects of physiology and behaviour. D. Phil thesis, University of Oxford, UK. Simpson, C. L., Chyb, S. and Simpson, S. J. (1990). Changes in chemoreceptor sensitivity in relation to dietary selection by adult Locusta migratoria L. Entomol. Exp Appl. 56, 259–268. Simpson, S. J. (1992). Mechanosensitive neurones in the suboesophageal ganglion of the locust. Physiol. Entomol. 17, 351–369.
THE NEUROBIOLOGY OF TASTE IN INSECTS
203
Simpson, S. J. (1994). Experimental support for a model in which innate taste responses contribute to regulation of salt intake by nymphs of Locusta migratoria. J. Insect Physiol. 40, 555–559. Simpson, S. J. and Raubenheimer, D. (1993a). The central role of the haemolymph in the regulation of nutrient intake in insects. Physiol. Entomol. 18, 395–403. Simpson, S. J. and Raubenheimer, D. (1993b). A multi-level analysis of feeding behaviour: the geometry of nutritional decisions. Phil. Trans. R. Soc. Lond. B 342, 381–402. Simpson, S. J. and Raubenheimer, D. (1996). Feeding behaviour, sensory physiology and nutrient feedback: a unifying model. Entomol. Exp. Appl. 80, 55–64. Simpson, S. J. and Raubenheimer, D. (2000). The hungry locust. Advances in the study of behavior. 29, 1–44. Simpson, S. J. and Simpson, C. L. (1992). Mechanisms controlling modulation by amino acids of gustatory responsiveness in the locust. J. Exp. Biol. 168, 269–287. Simpson, S. J. and White, P. R. (1990). Associative learning and locust feeding: evidence for a ‘learned hunger’ for protein. Anim. Behav. 40, 506–513. Simpson, S. J., Simmonds, M. S. J., Wheatley, A. R. and Bernays, E. A. (1988). The control of meal termination in the locust. Anim. Behav. 36, 1216–1227. Simpson, S. J., James, S., Simmonds, M. S. J. and Blaney, W. M. (1991). Variation in chemo-sensitivity and the control of dietary selection behaviour in the locust. Appetite 17, 141–154. Slifer, E. H. (1954). The reaction of a grasshopper to an odorous material held near one of its feet (Orthoptera: Acrididae). Proc. R. Ent. Soc. Lond. [A] 29, 177–179. Slifer, E. H. (1956). The response of a grasshopper, Romalea microptera (Beauvois), to strong odours following amputation of the metathoracic leg at different levels. Proc. R. Ent. Soc. Lond. [A] 31, 95–98. Smith, B. H. (1991). The olfactory memory of the honey bee Apis mellifera. I. Odorant modulation of short- and intermediate-term memory after single-trial conditioning. J. Exp. Biol. 161, 367–382. Smith, B. H. and Getz, W. M. (1994). Nonpheromonal olfactory processing in insects. Annu. Rev. Entomol. 39, 351–375. Spector, A. C. (2000). Linking gustatory neurobiology to behaviour in vertebrates. Neurosci. Biobehav. Rev. 24, 391–416. Sta¨dler, E. (1986). Oviposition and feeding stimuli in leaf surface waxes. In: Insects and the Plant Surface (eds. Juniper, B., and Southwood, T. R. E.), pp. 105–122. London, U.K: Edward Arnold. Sta¨dler, E. and Hanson, F. E. (1975). Olfactory capabilities of the ‘gustatory’ chemoreceptors of the tobacco hornworm larvae. J. Comp. Physiol. 104, 97–102. Stocker, R. F. (1994). The organization of the chemosensory system in Drosophila melanogaster – a review. Cell Tissue Res. 275, 3–26. Strausfeld, N. J. (1976). Atlas of an Insect Brain. Berlin – Heidelberg – New York: Springer. Szentesi, A. and Jermy, T. (1990). The role of experience in host plant choice by phytopagous insects. In: Insect-Plant Interactions, Vol. 2. (ed. Bernays, E. A.), pp. 39–74. Boca Raton, Florida: CRC press. Takken, W., van Loon, J. J. A. and Adam, W. (2001). Inhibition of host-seeking response and olfactory responsiveness in Anopheles gambiae following blood feeding. J Insect Physiol. 47, 303–310. Tousson, E. and Hustert, R. (2000). Central projections from contact-chemoreceptors of the locust ovipositor and adjacent cuticle. Cell Tissue Res. 302, 285–294.
204
S. M. ROGERS AND P. L. NEWLAND
Thompson, K. J. (1986a). Oviposition digging in the grasshopper. I. Functional anatomy and the motor programme. J. Exp. Biol. 122, 387–411. Thompson, K. J. (1986b). Oviposition digging in the grasshopper. II. Descending neural control. J. Exp. Biol. 122, 413–425. Tully, T. (1987). Drosophila learning and memory revisited. Trends Neurosci. 10, 330–335. van Loon, J. J. A. (1990). Chemoreception of phenolic acids and flavonoids in larvae of two species of Pieris. J. Comp. Physiol. A 166, 889–899. van Loon, J. J. A. (1996). Chemosensory of feeding and oviposition behaviour in herbivorous insects: a glance at the periphery. Entomol. Exp. Appl. 80, 7–13. Varanka, I. (1981). Taste reception by maxillary palps of the migratory locust. Advances in Physiological Science 23, 459–478. Vickers, N. J., Christensen, T. A. and Hildebrand, J. G. (1998). Combinatorial odor discrimination in the brain: attractive and antagonist odor blends are represented in distinct combinations of uniquely identifiable glomeruli. J. Comp. Neurol. 400, 25–56. Visser, J. H. (1986). Host odor perception in phytophagous insects. Ann. Rev. Entomol. 31, 121–144. Wacht, S., Lunau, K. and Hansen, K. (1996). Optical and chemical stimuli control pollen feeding in the hoverfly Eristalis tenax. Entomol. Exp. Appl. 80, 50–53. Watson, A. H. D. and Burrows, M. (1985). The distribution of synapses on the two fields of neurites of spiking local interneurones in the locust. J. Comp. Neurol. 240, 219–232. White, P. R. and Chapman, R. F. (1990). Tarsal chemoreception in the polyphagous grasshopper Schistocerca americana: behavioural assays, sensilla distributions and electrophysiology. Physiol. Entomol. 15, 105–121. White, P. R., Chapman, R. F. and Ascoli-Christensen, A. (1990). Interactions between two neurones in contact-chemosensilla of the grasshopper Schistocerca americana. J. Comp. Physiol. A 167, 431–436. Wibe, A., Borg-Karlson, A.-K., Norin, T. and Mustaparta, H. (1996). Identification of plant volatiles activating the same receptor neurones in the pine weevil, Hylobius abietis. Entomol. Exp. Appl. 80, 39–42. Woodhead, S. and Chapman, R. F. (1986). Insect behaviour and the chemistry of plant surface waxes. In: Insects and the Plant Surface (eds. Juniper, B., and Southwood, T. R. E.), pp. 123–135. London, U.K: Edward Arnold. Woodrow, D. F. (1965). The responses of the African migratory locust Locusta migratoria migratorioides R and F to the chemical composition of the soil at Oviposition. Anim. Behav. 13, 348–356. Yetman, S. and Pollack, G. S. (1986). Central projections of labellar taste hairs in the blowfly, Phormia regina Meigen. Cell Tissue Res. 245, 555–561. Yoshida, M. and Kobayashi, M. (1992). Identified neurones involved in the control of rhythmic buccal motor activity in the snail Achatina fulica. J. Exp. Biol. 164, 117–133. Zacharuk, R. Y. (1980). Ultrastructure and function of insect chemosensilla. Ann. Rev. Entomol. 25, 27–47. Zacharuk, R. Y. (1985). Antennae and Sensilla. In: Comprehensive Insect Physiology, Biochemistry, and Pharmacology, Vol. 6. (ed. Kerkut, G. A., and Gilbert, L. I.), pp. 1–69. Oxford: Pergamon.
Trehalose – The Insect ‘Blood’ Sugar S. Nelson Thompson Analytical Chemistry Instrumentation Facility and Department of Entomology, University of California, Riverside, CA 92521, USA
1 Introduction 206 1.1 Discovery of trehalose 206 1.2 The insect ‘blood’ sugar 207 1.3 Homeostasis and enantiostasis 208 1.4 Non-homeostatic regulation and trehalose function 209 2 Chemistry 210 2.1 Properties 210 2.2 Analysis 213 3 Occurrence with other haemolymph metabolites 214 4 Metabolism 220 4.1 Biosynthesis 220 4.2 Degradation 234 5 Hormonal regulation of metabolism 241 5.1 Hypertrehalosemic factors 241 5.2 Hypotrehalosemic factors 245 5.3 Hormonal regulation of polyol synthesis during cold-hardening 5.4 Diapause hormone and trehalose hydrolysis 247 6 Interactions of trehalose and lipid metabolism 248 7 Physiological roles 248 7.1 Energy storage 248 7.2 Stress protection 249 7.3 Regulation of feeding 253 8 Conclusion 259 Acknowledgements 261 References 261
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Abstract Trehalose, the non-reducing disaccharide of glucose, is the principal sugar circulating in the blood or haemolymph of most insects. Resistance to acid hydrolysis and an absence of direct intramolecular hydrogen bonding make trehalose chemically unique when compared with other common disaccharides, ADVANCES IN INSECT PHYSIOLOGY VOL. 31 ISBN 0-12-024231-1 DOI: 10.1016/S0065-2806(03)31004-5
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particularly sucrose, the non-reducing disaccharide of plant origin. Synthesized in the fat body following digestion of dietary sugar, trehalose is a condensation product of two glycolytic intermediates, glucose-1-phosphate and glucose6-phosphate. Alternative sources of trehalose are glycogen breakdown and gluconeogenesis. Hydrolysis to reform glucose, catalyzed by isozymes of a single enzyme, trehalase, is the only known pathway of trehalose utilization. Trehalose synthesis and degradation are under hormonal control involving both hypertrehalosemic and hypotrehalosemic factors. Trehalose concentration in the blood, however, is not homeostatically regulated. Rather, trehalose occurs at highly variable levels, typically between 5 and 50 mM, depending on environmental conditions, physiological state and nutrition. This variable concentration is essential for fulfilling the roles of trehalose, as (1) an energy store, the traditional role ascribed to trehalose; (2) a cryoprotectant, reducing the supercooling point of some freeze-avoiding insects; (3) a protein stabilizer during osmotic and thermal stress, a function only recently investigated in insects, and (4) a component of a feedback mechanism regulating feeding behaviour and nutrient intake, where blood metabolite levels including trehalose act through modulation of taste receptor responses and through the central nervous system to influence food selection. These are all examples of functional conservation in the absence of homeostasis. This has been termed enantiostasis, where functional conservation serves as a mechanism of physiological adaptation despite what appears to be an unstable internal milieu.
1 1.1
Introduction DISCOVERY OF TREHALOSE
Trehalose was isolated in 1832 from rye ergot (Wiggers, 1832). The sugar was first known as mycose. Subsequently described and renamed by Berthelot (1858) from the weevil genus Larinus, trehalose is the principal sugar in the blood of insects. The name refers to the beetle’s pupal cells or cocoons – ‘trehala’ – formed around the stems of host plants collected by people throughout Asia Minor (Pierce, 1915). The trehala, also known as tricara or tricala, are widely used as a sweet food and as a medicine. Medicinal applications are described in Persian literature dating from the mid-seventeenth century (Hanbury, 1859). Trehalose has since been described from diverse sources including bacteria, fungi, lichens, algae, a few vascular plants, and various invertebrate animals. Ironically, trehalose, produced by genetically modified plants, is being commercialized as a superior artificial sweetener (Kidd and Devorak, 1994; Portmann and Birch, 1995). Wyatt (1967) provides a detailed history of the finding and various historic uses of trehalose.
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Interest about trehalose in insects followed its simultaneous discovery by Wyatt and Kalf (1956) in pupae of the silkmoth Antheraea polyphemus, and by Howden and Kilby (1956) in Schistocerca gregaria, the desert locust. As the principal circulating carbohydrate of insects, trehalose is notable because of the high concentrations at which it often occurs, but perhaps more important, is its variable concentration. Trehalose concentrations in insect blood are typically greater by an order of magnitude than the levels of glucose found in the blood of mammals and other vertebrate animals. Trehalose, however, is only one of many metabolites occurring at high concentrations in insect blood. Glucose is often present, sometimes at levels similar to those of mammalian blood, but generally lower. Amino acids, proteins and other organic compounds are also present, and their concentrations may exceed that of trehalose (Chen, 1985; Mullins, 1985). Strictly speaking, the circulating fluid of insects and other invertebrates is dissimilar from the blood of vertebrate animals in that it is not contained within a closed system (Miller, 1985, 1997). Due to the absence or rudimentary presence of a mesoderm-derived lining of the body cavity the circulating fluid directly contacts the insect tissues and organs, and is therefore more akin to mammalian lymph. Hence the term haemolymph, the proper name for insect blood. Haemolymph circulates within a haemocoel, rather than a coelom or true body cavity. The haemocoel forms during embryonic development following confluence of the coelomic sacs with the epineural sinus (Chapman, 1998). Among organs and tissues, the embryonic mesoderm ultimately forms the fat body, the metabolically active organ or tissue lying within the haemocoel, the dorsal vessel and the circulating cells or haemocytes (Mori, 1979). Open circulation is not random. Haemolymph flow through the sinuses is largely unidirectional and facilitated by a contractile dorsal vessel, as well as segmental vessels and/or pulsatile organs that direct haemolymph throughout the haemocoel and into the extremities (Miller, 1985; Pass, 2000). Muscle contractions and ventilation also assist in circulating the haemolymph (Miller, 1997; Wasserthal, 1997). Direct contact of the haemolymph with the internal organs facilitates the diffusion of metabolites and other compounds from the organs and tissues into the haemolymph, perhaps explaining their high levels. For the same reason, mammalian lymph, also in contact with tissues and organs, would be expected to have high levels of circulating metabolites, but does not. A feature distinguishing insect haemolymph, is the presence within the haemocoel of the gut, from which the products of digestion are absorbed directly into the haemolymph and the open circulation. The high concentration of circulating metabolites and nutrients characteristic of open circulation is often considered an accommodation for inefficient circulation (Friedman, 1985; Mullins, 1985). Certainly, high metabolite and nutrient levels in open circulatory systems provide the distinct advantage of
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rapid substrate availability to tissues in support of energy production and other physiological functions. However, the small size of even large insects negates the need for a high-pressure closed system to transport fluid over long distances. Mixing time, the time required for the complete and uniform dispersal of circulating compounds throughout the haemocoel, is highly variable, but may be as little as a few minutes (Coon, 1944; Cherbas and Cherbas, 1970; Kovalova and Strunecka, 1973). Mixing time depends on an insect’s activity, stage of development, and the exact circulatory dynamics involved (Jones, 1977). The transport of sugar and other energy substrates that maintain the exceedingly high rates of aerobic respiration typical of flight by many adult insects (see Section 4.2.2.2) attests to the circulatory efficiency of insects. Thus, open circulation is not synonymous with inefficient circulation. Availability of O2 is not a consideration as insects have a highly efficient tracheal system for transporting O2 directly to tissues. The high concentrations of haemolymph metabolites and nutrients, and particularly the variability of concentrations, are essential for basic physiological functions not directly related to circulation. 1.3
HOMEOSTASIS AND ENANTIOSTASIS
Past reviews of trehalose and its metabolism in insects (Friedman, 1978; Jungreis, 1980; Mullins, 1985; Nijhout, 1994; Becker et al. 1996) have emphasized the homeostatic regulation of haemolymph trehalose, often citing Claude Bernard’s principle, ‘la fixite du milieu interieur’ (Bernard, 1865). This central physiological concept was elaborated by Cannon (1932) as ‘homeostasis’, in reference to the stable composition of body fluids in mammals and the complex feedback mechanisms involved in maintaining the internal environment within narrow limits. Trehalose and other circulating organic metabolites, however, are not maintained within narrow limits in insect haemolymph. Trehalose occurs at variable levels depending upon physiological state and nutritional conditions. Moreover, trehalose concentrations vary between individuals of the same developmental stage of the same insect species. Hence, trehalose is not regulated in a homeostatic manner, and the principal functions of trehalose in the physiology of insects necessitate its presence at variable concentration. That is not to say that haemolymph trehalose level and trehalose metabolism are unregulated, but rather that the emphasis is on functional regulation rather than the regulation of state. Many cellular and metabolic processes in invertebrates as well as vertebrates involve strategies of adaptation where homeostasis is not achieved (Boiteux et al., 1980; Hochachka and Somero, 1984). The concept of functional conservation as a mechanism for physiological adaptation was formally termed enantiostasis by Mangum and Towle (1977). ‘An enantiostatic regulation occurs when the effect of change in one chemical or physical property of the internal milieu is opposed by a change in another’.
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209
Mangum et al. (1976) described enantiostatic regulation of osmotic pressure and hypoxic tolerance in the estuarine blue crab Callinectes sapidus. On migrating upstream from salt to fresh water, crabs are exposed to a severely oxygen-depleted environment and at the same time display a dramatic decrease in haemolymph osmotic concentration. Intracellular osmotic equilibrium is achieved, not by loss of salts, but by a decrease in the concentration of intracellular free amino acids by deamination. The resultant release of ammonia produces a sharp increase in haemolymph pH, directly proportional to haemolymph osmotic concentration. This change in haemolymph pH causes a shift in the oxygen dissociation curve for haemocyanin that enables efficient oxygen release at lower salinity, enabling the crab to maintain aerobic respiration in the hypoxic environment. Thus, during enantiostasis, the internal milieu may be unstable, but the net result of regulation on a particular physiological system is stability. Similar investigations demonstrating functional conservation due to the opposing actions of physiological and/or metabolic processes in insects are lacking, but future studies will undoubtedly demonstrate that enantiostasis plays an important role in physiological regulation by insects. The principal role ascribed to trehalose in insects by most authors is an energy store, a function consistent with Mellanby’s (1939) original view of the haemolymph as a reserve. Haemolymph trehalose clearly serves this end, as numerous studies on insect flight and flight muscle metabolism demonstrate (Section 4.2.2.2). That function, however, might be argued for many circulating metabolites, regardless of concentration, as it can be for blood glucose in mammals. Unlike those molecules generally considered storage metabolites, fat and glycogen, trehalose does not occur in a polymerized insoluble form, nor is it confined to an intracellular compartment. The view that variable trehalose concentration, as well as that of other circulating metabolites, serves a greater role in physiological function is not new. Jungreis (1980) described the haemolymph as a ‘dynamic’ tissue, and dismissed the homeostatic view of haemolymph as a mere reserve or ‘sink’ for metabolites. Regarding trehalose, Jungreis estimated that the steady-state turnover rate in resting larvae of the tobacco hornworm Manduca sexta are much higher than the rates for storage metabolites, and not unlike those of circulating mammalian blood metabolites when the total size of the trehalose pool is considered. Trehalose rapidly exchanges between haemolymph and fat body. 1.4
NON-HOMEOSTATIC REGULATION AND TREHALOSE FUNCTION
Among the functions of trehalose that necessitate its non-homeostatic regulation are: (1) Cryoprotection. Trehalose and other small metabolites accumulate to lower the supercooling point of haemolymph, allowing insects to avoid freezing at subzero temperatures. Alternately, due to special
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S. N. THOMPSON
non-colligative properties, trehalose enables insects to tolerate extracellular freezing and protects cells from resultant osmotic stress and dehydration. (2) Stress protection. It is proposed here that trehalose probably also protects insects from the deleterious effects of osmotic, anoxic and other environmental stresses. (3) Regulation of nutrient intake. Numerous investigations demonstrate that haemolymph levels of metabolites, including trehalose, provide an index of an insect’s nutritional status, and serve to regulate nutrient intake and feeding behaviour through physiological feedback mechanisms. Below, the author reviews the occurrence and metabolism of trehalose in insects, the hormonal regulation of metabolism and the principal physiological roles of the insect blood sugar. Such a review inevitably raises the question: Why is trehalose, rather than another disaccharide, sucrose for example, the blood sugar of insects? The answers lie in the chemical properties of the molecule.
2 2.1
Chemistry PROPERTIES
Trehalose, characterized by Maquenne (1891), is a disaccharide of D-glucose in which the anomeric carbons form the glycosidic linkage (Fig. 1). The empirical formula is C12H22O11. 1-D-glucopyranosyl- 1-D-glucopyranoside, -trehalose, is the only isomer found in nature, although (isotrehalose) and (neotrehalose) isomers have been synthesized and characterized (Birch, 1963; Dowd and Reilly, 1992). Trehalose is highly soluble in water and aqueous ethanol and crystallizes as a dihydrate with a melting point of approximately 96 C. The water of crystallization escapes at about 130 C and anhydrous trehalose melts at 203 C. Trehalose, like sucrose, the disaccharide of plant origin, is a non-reducing sugar. Both trehalose and sucrose fail simple Fehling’s and Benedict’s tests for oxidizing the free aldehyde and ketones functional groups formed by mutarotation in reducing aldoses and ketose such as glucose, maltose or fructose. Due to limited reactivity compared with common reducing sugars, non-reducing sugars have been termed ‘less toxic’ (Candy et al., 1997) and ‘compatible’ (Behm, 1997). This allows for the presence of high stable concentrations in biological fluids, trehalose in insect haemolymph and sucrose in the vascular fluid of plants. In contrast to sucrose, 1-D-glucopyranosyl- 2-Dfructofuranoside, however, trehalose is much more resistant to acid hydrolysis, a characteristic consistent with the sole presence of pyranose moieties in trehalose (Moelwyn-Hughes, 1930). The rate constant for the acid hydrolysis of sucrose is K ¼ 14,600 106, and for trehalose, K ¼ 0.864 106 (Moelwyn-Hughes, 1929). These correspond to activation energies of 25.83 kcal/mole for sucrose and 40.18 kcal/mole for trehalose. The difference may be due to the greater reduction in
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211
FIG. 1 Structure of -trehalose. upper pyranose conformation, lower threedimensional crystal structure of the dihydrate. (After Liu et al., 1997.)
internal strain that occurs during formation of the five member furanose ring in sucrose compared with the six-member pyranose ring in trehalose (Eliel, 1962). The resistance of trehalose to acid hydrolysis, more than any other property, may explain why trehalose, rather than sucrose, is the haemolymph sugar of insects. Insect haemolymph typically contains high levels of proteins, peptides and free amino acids. Were trehalose to be readily hydrolyzed by the amine and free amino groups of these metabolites, subsequent Maillard reactions, condensation reactions between the aldehyde and primary and secondary amines, could lead to formation of imines and enamines, respectively, changing chemical properties and potentially causing deleterious structural modifications to proteins. An analogous situation is the ‘browning’ that occurs during food preservation, where the above reactions ultimately lead to the formation of pigments, and where trehalose is more effective than sucrose in preventing protein degradation (Rosser, 1991; O’Brien, 1996). Likewise, this may explain why most insects have low levels of haemolymph glucose. Regarding vascular plants, phloem saps often have sucrose concentrations in the range of 80–100 mM, comparable with trehalose in insect haemolymph, although they may have significantly higher concentrations (see A. E. Douglas, this volume). Proteins
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and amino acids, however, are generally at very low levels (Dale and Sutcliffe, 1986; Taiz and Zeiger, 1998). Trehalose, also unlike sucrose and other common disaccharides, lacks direct intramolecular hydrogen bonding between the hydroxyl groups of the pyranose moieties. However, in trehalose dihydrate the two glucose units are linked indirectly by hydrogen bonding with the two water molecules (Taga et al., 1972). One water molecule links O2 and to O0 4, while the second links O2 and O0 6 (Fig. 2). Conformational analysis demonstrates that trehalose dihydrate has a minimum energy region comprised principally of a single well where both pyranose rings have bond angles about the glycosidic oxygen of approximately 44 (Dowd and Reilly, 1992). Neither the ring, nor the glycosidic oxygens are involved in hydrogen bonding but all hydroxyl groups are hydrogen bonded with water. In the crystalline state, hydrogen bonding produces long linear chains of trehalose dihydrate. Many sugars vitrify upon dehydration to form glassy amorphous solids where the arrangement of molecules and pattern of hydrogen bonding between molecules is highly unordered (Green and Angell, 1989; Aldous et al., 1995). Among sugars, trehalose displays a higher glass transition temperature, regardless of water content to form highly stable glasses that are not hygroscopic (Levine and Slade, 1992; Crowe et al., 1996; Wang and Haymet, 1998). Glasses formed by most other sugars, including sucrose, slowly absorb water, and crystallize.
FIG. 2 Hydrogen bonding in -trehalose dihydrate. Molecule is shown perpendicular to the approximate two-fold axis. (After Taga et al., 1972.)
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Many investigators believe that the collective physical and chemical properties of trehalose make this disaccharide uniquely suitable as a general stress protector and preservative (see Section 7.2). 2.2
ANALYSIS
Numerous methods are available for isolation and qualitative and quantitative analysis of trehalose and other sugars in animal and plant tissues. An in-depth discussion of the methodology is beyond the scope of this article. Only a brief overview of the common methods used for estimating trehalose in insect and other biological materials is presented here (Table 1 – with selected references). A simple method employed in many insect studies cited below, is the spectrophotometric analysis of haemolymph following deproteinization and treatment with anthrone reagent. If trehalose is the major non-glycogen carbohydrate present, this technique provides a crude but approximate measure of trehalose concentration. Any contribution from glucose can be accounted for by applying one of the well-known enzymatic/ spectrophotometric assays for glucose, and subtracting this result from that of the anthrone analysis of total sugar. Alternatively, samples can be analyzed for glucose before and after treatment with trehalase or alkaline hydrolysis. The difference between the two results reflects the concentration of trehalose. A modification of this method employs flow injection analysis with immobilized
TABLE 1
Methods for analysis of sugars in insect haemolymph and other tissues
Method Spectrophotometric assay/anthrone reagent Flow injection analysis/trehalase
Reference
van Handel, 1965; Kramer et al., 1978; Ferreira et al., 1997 Schulze et al., 1995; Meyer zu Duttingdorf et al., 1997 Thin layer chromatography Zlatkis and Kaiser, 1977; Fell, 1990; (TLC)/high performance TLC Fried and Sherma, 1999 Gas-liquid chromatography Sweeley et al., 1963; Knapp, 1979; Hawakawa and Chino, 1982a; Abou-Seif et al., 1993; Grob, 1995 High-pressure liquid chromatography Honda, 1984; Iida and Kajiwara, 1990; (HPLC) Ga¨de, 1991; La Course, 1995; Ferreira et al., 1997; Hallsworth and Magan, 1997 Ion exchange HPLC Murray et al., 1997 1 H Nuclear magnetic resonance Thompson, 1990; Fan, 1996; spectroscopy (NMR) Phalaraksh et al., 1999 13 C NMR Coxon, 1980; Bock and Pedersen, 1983; Kukal et al., 1988; Thompson, 1990, 1998
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S. N. THOMPSON
trehalase. If trehalose is not the principal sugar present in a sample, extraction and isolation of total sugars, for example by ethanol extraction, will be necessary, followed by a more thorough analysis. In the case of samples whose chemical composition is unknown, preliminary analysis is required, after which the suitability of using less exacting methods in further investigation can be evaluated. Paper and thin layer chromatography can be effective tools for analysis of sugars, but are not widely employed because of limited sensitivity. Some success, however, has been achieved with high-performance thin layer chromatography (HPTLC) for separation and semi-quantitative analysis of sugars, including trehalose. Gas/liquid chromatography following chemical derivatization of sugars is quantitative, reasonably sensitive, and widely employed. High-pressure liquid chromatography (HPLC) which does not require derivatization and is highly sensitive is now the preferred method. All chromatographic methods require sugar extraction and some degree of purification. The necessity for these preliminary procedures may be significantly reduced by use of more specialized HPLC applications, for example ion exchange HPLC. Proton and 13C nuclear magnetic resonance spectroscopy (NMR) have proven valuable methods for quantitative analysis of sugars. The method has the distinct advantage that all sugars, as well as all other metabolites, are detected simultaneously. Analysis of biological fluids, however, may result in complex spectra where the individual components are not easily resolvable, in which case some extraction and isolation procedures may be required. The 13C spectrum of trehalose has six distinct signals for the combined carbons of the individual glucose units, and is simple by comparison with free glucose and most other sugars. NMR is quantitative but inherently insensitive. Many specialized applications of NMR are available for overcoming this limitation under a variety of circumstances, but quantification of metabolites present at low levels will often require additional analysis employing methods that are more sensitive.
3
Occurrence with other haemolymph metabolites
The low level of circulating glucose or reducing sugar in insects was recognized before the last century. By mid-century, it was apparent that trehalose was a principal circulating form of glucose in insects. With the exception of flight muscle, trehalose levels in tissues other than haemolymph are suspect, owing to the difficulty of eliminating contamination by haemolymph. Several reviews provide lists of trehalose concentrations for numerous insect species (Wyatt, 1967; Florkin and Jeuniaux, 1974; Bedford, 1977; Kramer et al., 1978; Mullins, 1985). These demonstrate that the static haemolymph trehalose concentration
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215
of insects maintained under a wide variety of conditions generally varies between 10 and 50 mM. Lower levels are common, and the highest occur in lepidopteran and coleopteran insects where trehalose often is >100 mM. In contrast, glucose is seldom present at concentrations above 5 mM and is often below 1 mM. Exceptions occur among dipteran, and particularly hymenopteran species. Often in these insects, haemolymph trehalose and glucose are at moderate to high concentrations, with glucose sometimes higher than trehalose. Larvae of the blow fly, Phormia regina, for instance, are reported to have haemolymph glucose in the range of 5–10 mM and only traces of trehalose (Evans and Dethier, 1957; Wimer, 1969). Adults have glucose levels as high as 30 mM with trehalose approaching 50 mM (Evans and Dethier, 1957) (Table 2). Calliphora erythrocephala has approximate haemolymph concentrations for glucose and trehalose of 75 and 50 mM, respectively (Duve, 1978) (Table 2). Honeybee workers, Apis mellifera, have been reported with haemolymph glucose as high as 75 mM and trehalose 40 mM (Leta et al., 1996), but other studies report levels half as great (Arslan et al., 1986). Considerable differences in sugar concentrations are reported between bee strains, with trehalose often higher than glucose (Bounias, 1981). In queens, concentrations of approximately 50 mM glucose and 32 mM trehalose have been found (Alumot et al., 1969). Other sugars, particularly fructose, are also present in the haemolymph of the honeybee, reflecting the high sucrose content of nectar (Bounias and Morgan, 1984). Although there has been some question concerning the accuracy of sugar concentrations reported for the honeybee, as the haemolymph has a high activity of glycosidase (Abou-Seif et al., 1993), this may reflect the mechanisms regulating haemolymph sugar levels. Haemolymph sugar levels in the honeybee are related to metabolic rate and crop emptying (Blatt and Roces, 2001). When the crop is filled with sucrose solution, the honeybee displays equivalent haemolymph levels of trehalose, glucose and fructose at metabolic rates up to approximately 5 ml CO2/h (Table 2). At higher metabolic rates, trehalose concentration decreases, while glucose and fructose levels increase until depletion of sucrose from the crop. At that point, the rate of decrease in haemolymph trehalose increases and the levels of fructose and glucose remain nearly constant as metabolic rate increases. With little glycogen stores and an inability to use amino acids for energy, dietary sucrose passed through the proventriculis, and digested and absorbed from the gut, is the principal source of energy for the honeybee. Thus, there is a linear relationship between metabolic rate and the rate of sugar transport through the proventriculus (Blatt and Roces, 2002a). With an upper limit to the rate of fat body trehalose synthesis from absorbed sugar, crop emptying is important in determining the levels of haemolymph sugars. Haemolymph trehalose concentration is important in the feedback mechanism regulating the activity of the proventriculus (Blatt and Roces, 2002b).
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Studies with the face fly, Musca automnalis, demonstrate a clear circadian rhythm for haemolymph trehalose concentration (Hayes et al., 1990), a factor not considered in most investigations. In adult flies, trehalose concentration is at a high of approximately 70-mM at 16 h following lights on, after which lights were turned off. Trehalose then decreases to 11 mM, at 8 h after lights off. It should be apparent from the above discussion that haemolymph trehalose level is best understood as a dynamic component, made relevant by considering the metabolic, physiological and behavioural factors that influence and are influenced by it. Thus, haemolymph sugar levels in individual insects are not discussed further here. Rather, examples are provided in Table 2 to illustrate how physiological state and various experimental treatments affect haemolymph concentrations of trehalose and other sugars in select insect species. In addition to sugars, haemolymph contains high levels of other organic solutes, including lipids, proteins, lipoproteins, peptides and particularly amino acids (Wyatt and Pan, 1978; Chen, 1985; Mullins, 1985; Telfer and Kunkel, 1991). In most insects, the latter contribute significantly to total haemolymph osmolar concentration, more than any other organic component, including trehalose (Sutcliffe, 1963; Machin, 1981). As with carbohydrates, the qualitative and quantitative haemolymph composition of other organic compounds is highly variable between and within individual species and is markedly affected by developmental stage, nutritional status, physiological state, and so on. High haemolymph levels of nitrogenous compounds may seem paradoxical in some dipteran and hymenopteran species having significant levels of reducing sugars, if condensation reactions are prevalent and affect proper functioning and/or structural integrity of haemolymph peptides and proteins (see Section 2.1). Such effects, however, may be partially mitigated by pH. The rate of reaction for condensation of ketones and aldehydes with amines is highly sensitive to pH in the physiological range of insect haemolymph (Jencks, 1959). The precise pH for maximal rate will depend in part on the pKa for the amine and the presence of associated carbonyl groups. Moreover, in the case of primary amines, addition or elimination may be rate limiting depending on pH (Cordes and Jencks, 1962). With some haemolymph constituents, storage proteins for example (Telfer and Kunkel, 1991), the level in the haemolymph may be sufficiently high that any reaction with reducing components has a minimal overall impact. Other mechanisms likely also exist for masking or minimizing reactions involving reducing components. Investigations that simultaneously examine the complete organic composition of haemolymph in a single stage of any insect are lacking. It is, therefore, not possible to establish consistent and reliable correlations between the concentrations of various haemolymph components. Appropriately,
Haemolymph sugar (mM)
Diptera Calliphora erythrocephala adult
Phormia regina adult Dictyoptera Blaberus discoidalis adult Hymenoptera Apis mellifera foraging adult
Apis mellifica adult Apis mellifica worker adult
Condition
Reference
Trehalose
Glucose
45 ! 58 ! 32
88 ! 111 ! 95
24 ! 53
28 ! 11
20 ! 25
Not determined
15 min following injection of octopamine into decapitated insects
Park and Keeley, 1996
92 ! 92 ! 13
54 ! 54 ! 54
Blatt and Roces, 2001
92 ! 92 ! 50
28 ! 28 ! 80
105 ! 80 ! 50
28 ! 28 ! 80
90 ! 50 42 ! 21 88 ! 7
28 ! 8 28 ! 14 4!3
Effect of increasing metabolic rate (0.5 ! 5 ! 9 ml CO2/h) -after feeding on a 15% sucrose solution -after feeding on a 30% sucrose solution -after feeding on a 50% sucrose solution After 7 h starvation After walking 10 meters 3 h following injection of midgut insulin-like peptide
24 h following extirpation of median neurosecretory cells (MNC) and 4 h after injection of MNC homogenate. Flies were provided sucrose 48 h following glucose feeding
Duve, 1978
Evans and Dethier, 1957
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
TABLE 2 Effects of physiological condition and various experimental treatments on haemolymph trehalose and glucose concentrations in select insect species
Abou-Seif et al., 1993 Bounias et al., 1986
217
(Continued)
218
TABLE 2 Continued Haemolymph sugar (mM)
Lepidoptera Bombyx mori 5th instar larva
Helicoverpa zea 5th instar larva
Glucose
Condition
13 ! 14 ! 11 13 ! 8 ! 6
0.4 ! 0.2 ! 0.1 0.4 ! 0.2 ! 0.1
11 ! 4 11 ! 15
Not determined Not determined
Trace ! 4
Not determined
Starved after day 3, 0 ! 6 ! 12 h Neck ligated after day 3, 0 ! 6 ! 12 h 24 h after neck ligation 2 h following injection of brain/ corpora cardiaca/corpora allata extract Maintained on a carbohydrate free diet for 1 day after moulting and transferred to a high sucrose diet
4.0 ! 3.3
Not determined
4.0 ! 0.7
Not determined
35 ! 40 ! 35
1.5 ! 0.2 ! 0.2
13 ! 37
Not determined
40 ! 100
Not determined
Maintained on a high sucrose diet for 1 day after moulting and starved for 4 h Maintained on a high sucrose as above and transferred to a diet with a protein/carbohydrate ratio of 4:1 Starved after day 3, 0 ! 24 ! 48 h After 3 days feeding on a carbohydrate free diet with increasing amounts of casein After 3 days on diet with increasing amounts of sucrose
Reference Satake et al., 2000 Oda et al., 2000
Friedman et al., 1991
Siegert, 1987 Thompson et al., 2003
S. N. THOMPSON
Manduca sexta 5th instar larva
Trehalose
5 ! 0.1
After 48 h starvation of newly moulted unfed larvae 45 min following injection of corpora cardiaca extract
Dahlman, 1973
Pieris brassica 4th instar larva
22 ! 32
4!4
Philosamia Cynthia diapausing pupa
13 ! 105
Not determined
After 50 days storage at 20 C
Hawakawa and Chino, 1982b
105 ! 25
Not determined
9 days following transfer to 24 C after being maintained for 90 days at 2 C
Hawakawa and Chino, 1981
Odonata Tramea virginia larva
20 ! 50
Not determined
6 h following injection of cardiaca corpora extract
Tembhare and Andrew, 1991
Orthoptera Locusta migratoria adult
70 ! 35 mM
Not determined
30 min after initiation of flight
Gourdoux et al., 1989
van der Horst et al., 1978
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
72 ! 7
219
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contemporary studies focus on individual components and their physiological functions.
4
Metabolism
4.1 4.1.1
BIOSYNTHESIS
Biosynthesis from glucose
The 1– 1 glycosidic linkage of trehalose is formed by the trehalose phosphate synthase (EC 2.4.2.15)-catalysed condensation of glucose-6-phosphate with the glucose moiety of uridine diphosphoglucose (UDP-glucose) to form trehalose-6-phosphate in the fat body (Fig. 3). The reaction to form UDPglucose from glucose-1-phosphate is catalysed by glucose-1-phosphate uridylyl transferase (EC 2.7.7.9). The same reaction provides substrate for glycogen synthesis. Thus, trehalose is a product of two glycolytic intermediates. Although a few early studies suggested that trehalose synthesis occurred in
FIG. 3 Metabolic scheme illustrating the pathways for trehalose synthesis, glycolysis and gluconeogenesis.
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muscle and haemolymph, trehalose synthesis is now thought to occur exclusively in the fat body. A second reaction, catalysed by trehalose phosphatase (EC 3.1.3.12), hydrolyses the phosphate ester, followed by the release of trehalose into the haemolymph. First demonstrated in the desert locust, S. gregaria, by Candy and Kilby (1961), trehalose synthesis has since been demonstrated, principally from various 14C-enriched isotopomers of glucose, in fat body preparations of numerous insect species (Wyatt, 1967; Bailey, 1975; Friedman, 1985). Trehalose synthesis has also been characterized in noninsect organisms, including nematodes, yeast and other fungi, and plants such as Arabidopsis and some ferns (Elbein, 1974; Thevelein, 1984; van Laere, 1989; Panek and Panek, 1990; Muller et al., 1995; Behm, 1997; Goddijn and van Dun, 1999). Trehalose-6-phosphate synthetase has been partially characterized in fractionated fat body homogenates from Hyalophora cecropia silkmoth larvae (Murphy and Wyatt, 1965), as well as from fat body of adult blow fly, P. regina (Friedman, 1971) and Periplaneta americana, the American cockroach (Friedman and Hsueh, 1979). The enzyme from H. cecropia has an affinity or KM (Michaelis constant–substrate concentration for half-maximal velocity) of 0.3 mM for UDP-glucose and 5 mM for glucose-6-phosphate. Crude preparations of the enzyme display a sigmoid substrate–velocity relationship with glucose-6-phosphate, which stimulates trehalose phosphate synthase activity. On the other hand, trehalose inhibits the enzyme. The extent of inhibition depends not only on trehalose concentration but also on the concentrations of glucose-6-phosphate and Mg þ 2. In the absence of Mg þ 2, enzyme activity, indicated by the release of UDP from UDP-glucose, is almost completely inhibited at approximately 35 mM trehalose. Trehalose-6phosphatase is highly active in H. cecropia fat body and hydrolysis of trehalose-6-phosphate is not rate limiting. Early findings with H. cecropia suggest that trehalose synthesis and glycogen synthesis, catalysed by glycogen synthase (EC 2.4.1.11), are closely coordinated, because both syntheses utilize UDP-glucose as substrate. In H. cecropia fat body, soluble glycogen synthase has a KM ¼ 1.6 for UDPglucose, much lower than that of trehalose-6-phosphate synthase (Murphy and Wyatt, 1965). This suggests that trehalose synthesis predominates over glycogen synthesis at typically low concentrations of UDP-glucose. As trehalose concentration rises, however, trehalose, being an inhibitor of trehalose6-phosphate synthase, inhibits its own synthesis. This results in increased glucose-6-phosphate, which simultaneously activates glycogen synthase, inhibits glycogen phosphorylase, and increases glycogen synthesis. Coordination between trehalose and glycogen synthesis may also involve the kinetics of interconversion between different forms of glycogen synthase. Although little is known about the different forms of glycogen synthase in insects, the increased glycogen synthase activity in H. cecropia fat body in response to glucose-6phosphate was not accompanied by a change in KM for UDP-glucose. Thus,
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the enzyme examined by Murphy and Wyatt (1965) may be similar to the D (phosphorylated b) form in liver. Based on enzyme activity in the presence and absence of glucose-6-phosphate, the insect glycogen synthase preparation may be only 15–20% I (unphosphorylated a) form. In contrast, the soluble glycogen synthase of liver is mainly the I form, with a KM for UDP-glucose of 1.0 and 0.2 mM, in the absence and presence, respectively, of glucose6-phosphate. The lower value is near the KM ( ¼ 0.3) for UDP-glucose of trehalose-6-phosphate synthase from H. cecropia. The regulation of glycogen synthase in cold hardy lepidopteran larvae is discussed below (see Section 4.2.2.1) (Fig. 9). Trehalose-6-phosphate synthase in fat body of adult blow fly is dissimilar to the enzyme from H. cecropia, having a much higher affinity, KM ¼ 4.2 mM, for UDP-glucose and a lower sensitivity to inhibition by trehalose (Friedman, 1985). Like crude preparations of H. cecropia fat body, the partially purified enzyme from P. regina shows a sigmoidal substrate–velocity relationship with glucose-6-phosphate. Studies with trehalose-6-phosphatase purified from homogenates of whole insects, demonstrated that the Mg2 þ -dependent enzyme catalyses the hydrolysis of trehalose-6-phosphate (KM ¼ 0.6 mM), as well as glucose-6-phosphate (KM ¼ 0.29 mM) (Friedman, 1971). Clearly, the enzyme has a much higher affinity for the latter substrate. Glucose-6-phosphate and Pi are both competitive inhibitors of trehalose-6-phosphate hydrolysis, with KI ¼ 0.4 mM and 1.6 mM, respectively, indicating that the hydrolytic sites overlap at the phosphate binding site. Trehalose activates glucose-6-phosphate hydrolysis, increasing product formation and the affinity for glucose-6phosphate from KM ¼ 0.29 mM to KM ¼ 1.25 mM with maximal activation. Friedman and Hsueh (1979) compared the stimulation of glucose-6-phosphate hydrolysis by trehalose in preparations from 24 insect species representing six orders. They observed that activation of hydrolysis by trehalose was generally restricted to a select group of dipterans related to the family Calliphoridae. Regarding glycogen synthesis, the situation in P. regina fat body appears considerably different from that in H. cecropia, because the affinity of trehalose-6-phosphate for UDP-glucose, KM ¼ 4.2 mM, is higher, not lower, than that of glycogen synthase (KM ¼ 2.8 mM). Interestingly, P. regina and its close relatives, both as adults and larvae, are among those insects that sometimes display haemolymph levels of glucose that are equal to or greater than those of trehalose (see Section 3). Moreover, some adult dipterans, including P. regina, utilize proline in addition to sugar for flight fuel, and may have haemolymph proline levels that approach those of sugar (Bursell, 1981). It is perhaps not surprising then that glucose/trehalose/glycogen metabolic interactions differ between P. regina and H. cecropia. Trehalose-6-phosphate synthase from American cockroach fat body displays a KM ¼ 0.65 mM for UDP-glucose and for glucose-6-phospate, KM ¼ 5.5 (Friedman, 1985), similar to that of the enzyme from H. cecropia. In addition, like the H. cecropia enzyme, trehalose-6-phosphate synthase from P. americana
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
223
is highly sensitive to trehalose inhibition. Trehalose-6-phosphatase isolated from cockroach has a KM ¼ 2.09 mM for trehalose-6-phosphate, displays negligible affinity for glucose-6-phosphate, and is not activated by trehalose (Friedman and Hsueh, 1979). Extensive investigations have been conducted, principally with yeast and bacteria, on the molecular genetics of trehalose-6-phosphate synthase and trehalose-6-phosphatase (Winderickx et al., 1996; Wolschek and Kubicek, 1997; Goddijn and van Dun, 1999; Fillinger et al., 2001). Similar studies of the insect enzymes are lacking, but the trehalose phosphate synthase gene tps 1 has been cloned from Drosophila melanogaster (Chen et al., 2002). 4.1.2
Biosynthesis from dietary sugar
The natural foods of insects are extremely diverse (Slansky and Rodriguez, 1987). They vary in nutritional content from the high protein/low or no carbohydrate foods of carnivorous species, such as carrion and flesh-eating dipteran insects, to the low protein/high carbohydrate foods of aphids and other homopterans that feed on plant saps (Dadd, 1985). Published compilations describing the composition of artificial diets developed for laboratory rearing of a broad range of insects indicate that for optimal growth insects require a balance of digestible carbohydrate and protein (Singh, 1977; Reinecke, 1985). Phytophagous insects, feeding on plant material, often consume similar amounts of protein and digestible carbohydrate (Dadd, 1985), and have a rich complement of digestive proteases and carbohydrases with which to hydrolyse these to absorbable sugars and amino acids (Terra and Ferreira, 1994). Many insects feeding on extremely unbalanced diets have physiological adaptations in their alimentary tracts and/or have a robust compliment of gut symbionts, which may dramatically alter nutrient availability and utilization (Kaufman et al., 2000). In most cases, carbohydrate uptake is sufficient so that glucose for trehalose synthesis is derived mostly, if not entirely, from dietary sugar. Studies on sugar absorption in the midgut of the desert locust and the American cockroach establish that sugar uptake by passive facilitated diffusion is maintained by a glucose concentration gradient between the gut lumen and the haemolymph (Treherne, 1967). In the locust, 14C-enriched glucose, mannose and fructose are rapidly absorbed by the gastric caecae and subsequently metabolized to trehalose, presumably by the fat body. Passive absorption of sugar has also been demonstrated in several other species (Turunen, 1985). In some dipteran and hymenopteran insects, where haemolymph glucose level is relatively high, glucose transport may be facilitated by glycogen synthesis in parts of the midgut epithelium, or by other mechanisms of absorption including active transport (Dow, 1986). Investigations with larvae of the tobacco hornworm, M. sexta, maintained on a chemically defined diet, demonstrate a relationship between dietary sucrose
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FIG. 4 Haemolymph trehalose concentration in 5th instar Manduca sexta larvae feeding on a chemically defined artificial diet with variable levels of sucrose (left) and on a carbohydrate-free diet with variable levels of casein (right). The arrows signify the level of carbohydrate consumption below which larvae are gluconeogenic. (Data after Thompson et al., 2003.)
level, sucrose consumption and haemolymph trehalose concentration (Thompson and Redak, 2000; Thompson et al., 2001, 2002). On diets with a constant level of casein and variable sucrose, there is a positive relationship between sucrose consumption and haemolymph trehalose level (Thompson et al., 2003; Fig. 4 left). Under these conditions, there is no significant relationship between dietary protein level and haemolymph trehalose concentration, but in the absence of dietary sucrose there is a positive relationship between dietary protein level and trehalose concentration (Fig. 4 right). 4.1.3
Role of the pentose phosphate pathway in facilitating biosynthesis from dietary sugar
The pentose phosphate pathway competes with glycolysis in glucose degradation (Fig. 5). The pathway provides ribose sugar for nucleotide and nucleic acid synthesis and is a principal source of reduced NADPH þ H þ for reductive biosyntheses. During the oxidative phase, glucose-6-phosphate is decarboxylated to glucono--lactone-6-phosphate, catalysed by glucose-6-phosphate dehydrogenase (EC 1.1.1.49). After lactonase (EC 3.1.1.17)-catalysed conversion of glucono--lactone-6-phosphate to 6-phosphogluconate, further oxidation catalysed by 6-phosphogluconate dehydrogenase (EC 1.1.1.44) results in the formation of ribulose-5-phosphate and terminates the oxidative phase of the pathway (Fig. 5). NADP þ stimulates glucose-6-phosphate dehydrogenase and regulates this phase of the pathway. The fate of ribulose-5-phosphate depends on cellular demands. Where the demand is for nucleic acid biosynthesis, the products of the pathway are principally ribose-5-phosphate and reduced coenzyme, NADPH þ H þ , and the contribution of the second portion of the pathway, the non-oxidative phase, may be minimal. When cellular needs are for reducing power and energy, NADPH þ H þ and ATP, rather than ribose, ribulose-5-phosphate
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FIG. 5 Metabolic scheme illustrating the pentose phosphate pathway in relation to glycolysis.
formed in the oxidative phase is metabolized through the non-oxidative phase. The non-oxidative phase of the classical ‘F’ pathway involves phosphopentoisomerase (EC 5.3.1.6), phosphopentoepimerase (EC 5.1.3.1), transketolase (EC 2.2.1.1) and transaldolase (EC 2.2.1.2), enzymes catalysing the formation and condensation of ribose, xylulose, sedoheptulose and erythrose phosphates (Wood, 1985) (Fig. 5). If the primary cellular need is for reducing power, NADPH þ H þ , the final products are fructose-6-phosphate and glyceraldehyde-3-phosphate. Because the glucose phosphate isomerase (EC 5.3.1.9)catalysed reaction between fructose-6-phosphate and glucose-6-phosphate is generally at or near equilibrium, fructose-6-phosphate formed by the nonoxidative phase is metabolized to glucose-6-phosphate, and under these conditions the pathway forms a cycle. Glucose-6-phosphate so-formed may be recycled by the oxidative phase. On the other hand, cellular demand for ribose may exceed that for NADPH þ H þ . In that case, ribose demand may be fully met by reversal of the non-oxidative phase in the absence of ribose synthesis due to inhibition of the oxidative phase by NADPH þ H þ . The nonoxidative phase is fully reversible (Landau, 1985). In the case where energy production is the cellular priority, the final products of the non-oxidative phase, fructose-6-phosphate and glyceraldehyde-3-phosphate, are oxidized to pyruvate, via glycolysis. The actual operation of the pentose phosphate
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cycle is complex and dynamically balanced in a manner that simultaneously provides for all cellular demands. Most often, cellular need for NADPH þ H þ and ATP exceeds that for ribose sugars and to a varying degree ribose-5phosphate is metabolized to fructose-6-phosphate and metabolized by glycolysis. The pentose phosphate pathway of insects is incompletely understood. Several studies, conducted before 1980 and reviewed by Friedman (1985) and Ben Khay et al. (1987), demonstrated the presence of the pathway in several taxonomically diverse insects. These include a few dipteran species, larvae of the silkmoth Bombyx mori and cabbage butterfly Pieris brassicae, adult Tenebrio molitor, the migratory locust and the American cockroach. Most studies examined the formation of 14CO2 from various 14C-enriched isotopomers of glucose. One approach, first outlined by Wang et al. (1956), compares metabolism of (1-14C)glucose, (3,4-14C2) glucose or (3,4-14C) glucose and (6-14C)glucose. Via glycolysis, (1-14C)glucose and (6-14C)glucose are assumed to be oxidized to (3-14C)pyruvate. Metabolism by the pentose phosphate pathway only oxidizes (6-14C)glucose to (3-14C)pyruvate because (1-14C)glucose is decarboxylated in the first step of the oxidative phase with the 14 C released as 14CO2. 14CO2 formation from (3,4-14C2) glucose or (3,4-14C) glucose reflects aerobic respiration without distinguishing between the contributions of glycolysis and the pentose phosphate pathway. The measure of pentose phosphate pathway activity relative to glycolysis is calculated as the percent pentose pathway (Gp): Gp ¼ [G1 G6/G1 G6 þ G3,4] 100, where each component in the equation is the 14CO2 recovered from the isotopomer indicated. Based on this method, Horie et al. (1968) reported that the pentose phosphate pathway was responsible for 35% of overall glucose catabolism by intact B. mori larvae, while in the fat body the contribution was greater than 80%. Several recent studies on the pentose phosphate pathway have employed an even simpler measure, the ratio of 14CO2 obtained from (1-14C)glucose and (6-14C)glucose (Mtioui et al., 1993a,b, 1994, 1996; Alaoui et al., 1997), a ratio discussed by Katz and Wood (1960, 1963). Mtioui et al. and Alaoui et al. examined the effects of several factors, including a midgut insulin-like peptide, extracts of corpora cardiaca, adipokinetic hormone and a pyrethroid, deltamethrin, on pathway activity in L. migratoria and T. molitor. All the above investigations provide a qualitative measure of pentose phosphate pathway activity, and demonstrate that the pathway is very active in insects. At the same time, these studies are quantitatively inaccurate in estimating the relative contribution of the pathway to total glucose catabolism. The latter group of investigations assume that the sole product of the pathway is ribose sugar, and ignore any contribution of the non-oxidative phase. Studies employing the approach of Wang et al. assume that all fructose-6-phosphate and glyceraldehyde-3-phosphate produced by the non-oxidative phase are completely oxidized by glycolysis and the tricarboxylic acid cycle. These assumptions were untested and may be incorrect.
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Investigations employing in vivo NMR spectroscopy clearly demonstrate significant trehalose synthesis as a result of pentose cycling in M. sexta larvae (Thompson, 1999). Nevertheless, the pentose phosphate pathway facilitates the synthesis of trehalose because substrate metabolized through the pathway is selectively directed to pyruvate formation. When (2-13C)glucose is injected into larvae maintained on a high carbohydrate diet, the distribution of 13C in the carbon atoms of trehalose is consistent with the cycling of glucose through the pentose phosphate pathway to glucose-6-phosphate, prior to trehalose synthesis. The trehalose C1/C3 13C enrichment ratio is 2, in exact agreement with carbon rearrangements expected with the operation of the classical ‘F’ pentose phosphate cycle (Landau, 1985). This ratio remained constant as dietary sucrose level was decreased, but the relative level of pentose cycling, indicated by the trehalose C1/C2 13C enrichment ratio, decreased significantly. Employing (1,2-13C2)glucose as the administered substrate enables the contribution to trehalose synthesis from glucose metabolized directly via glycolysis to be distinguished from the contribution of glucose following passage through the pentose phosphate cycle (Fig. 6). When (1,2-13C2)glucose is incorporated directly into trehalose via glycolysis, a doublet (D) resonance signal is observed for both C1 and C2 of trehalose, due to spin–spin J coupling between the adjacent 13C-enriched carbon atoms. On the other hand, when (1,2-13C2)glucose is cycled through the pentose phosphate pathway to glucose-6-phosphate, the two 13C-enriched carbon atoms are uncoupled, giving rise to singlet (S) resonance signals for C1 and C3 of trehalose. The C1S/C2D 13C enrichment ratio is a relative measure of the two contributions. For insects maintained at the highest dietary carbohydrate level, the mean ratio is 0.22, demonstrating a significant but small contribution of pentose cycling to trehalose synthesis. The bulk of glucose for trehalose synthesis is derived directly from glycolysis. In a similar manner to that described above for trehalose, the contributions of glycolysis and the pentose phosphate pathway to pyruvate synthesis are estimated (Fig. 6). In this case, alanine synthesized by transamination of pyruvate is analysed. The measure of the two contributions is reflected by the C3S/C2D 13C enrichment ratio in alanine. The mean ratio is 0.41, significantly higher than the trehalose ratio above. If glucose-6-phosphate and fructose-6-phosphate were in equilibrium, the trehalose and the alanine ratios would be equivalent. The higher ratio in alanine suggests that under these nutritional conditions, the reaction is in disequilibrium and favours glucose-6-phosphate. Thus, the pentose phosphate pathway facilitates the synthesis of trehalose, because although significant cycling occurs, glucose metabolized through the pentose phosphate pathway to fructose-6-phosphate is preferentially oxidized to pyruvate over substrate metabolized directly from glucose via glycolysis.
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FIG. 6 Metabolic scheme illustrating the 13C enrichment of haemolymph trehalose and alanine synthesized from (1,2-13C2)glucose administered to 5th instar Manduca sexta larvae. The effects due to the pentose pathway and glycolysis are indicated, and the 13C multiplet signal structure expected from the 13C enrichment patterns or each metabolite are shown: D ¼ doublet signals due to J spin–spin coupling between adjacent 13 C atoms, S ¼ singlet signals. The numerals under the individual carbons refer to the derivation from the original administered (1,2-13C2)glucose. (After Thompson, 1999.)
4.1.4
Biosynthesis via gluconeogenesis
4.1.4.1 Gluconeogenesis from amino acids. Gluconeogenesis, the de novo or net synthesis of carbohydrate from non-carbohydrate precursors, principally amino acids, shares the reversible reactions of the glycolytic pathway (Fig. 3). The two metabolic processes are not a simple reversal of each other. Three steps are irreversible with the forward (glycolytic) and reverse (gluconeogenic) reactions catalysed by different enzymes, forming substrate cycles. The reactions of these substrate cycles provide for the short-term regulation of glycolytic/gluconeogenic flux as outlined by Pilkis and Claus (1991) for these pathways in liver. Carbohydrate synthesis from glucogenic amino acids begins
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in the mitochondrion with metabolism of alanine or other amino acids to pyruvate or oxaloacetate by transamination or via the tricarboxylic acid cycle. Pyruvate is carboxylated to form oxaloacetate, a reaction catalysed by pyruvate carboxylase (EC 6.4.1.1). After oxidation to malate, the latter metabolite is transported into the cytosol, where oxaloacetate is regenerated. Oxaloacetate then undergoes phosphoenolpyruvate carboxykinase (EC 4.1.1.32)-catalysed decarboxylation to phosphenolpyruvate. The substrate cycle formed, the pyruvate cycle, involves two enzymes in the gluconeogenic pathway, and a single enzyme, pyruvate kinase (EC 2.7.1.40) in the glycolytic pathway (Fig. 3). The second irreversible step forms the fructose phosphate cycle (see Section 4.1.6). During gluconeogenesis, condensation of the triose phosphates, glyceraldehyde-3-phosphate and dihydroxyacetone phosphate, forms fructose1,6-bisphosphate. The latter is dephosphorylated to fructose-6-phosphate, catalysed by fructose-1,6-bisphosphatase (EC 3.1.3.11). The corresponding forward reaction in the glycolytic pathway is catalysed by 6-phosphofructo-1kinase (EC 2.7.1.11). Finally, after isomerization of fructose-6-phosphate to glucose-6-phosphate, glucose is formed by glucose-6-phosphatase (EC 3.1.3.9)catalysed hydrolysis. In most insects, where haemolymph carbohydrate is principally trehalose, the glucose phosphate/glucose substrate cycle may be of lesser importance as a potential regulatory step than it is in animals with glucose as the blood sugar. Evidence for gluconeogenesis in insects was reported first by Wigglesworth (1942), from investigations of the mosquito Aedes aegypti. After storage reserves are exhausted by starvation, larvae fed alanine or glutamate display significant fat body glycogen deposition within 48 h. Authors of several investigations reviewed by Gourdoux et al. (1983) claim to observe gluconeogenesis, based on the incorporation of various 14C-enriched amino acids into glucose and other carbohydrates. A conclusion of net carbohydrate synthesis is suspect, however, because the contribution of the tricarboxylic acid cycle to glucose formation was not determined. Moreover, in most cases dietary conditions were unsuitable. In Tenebrio molitor for instance, Gourdoux et al. fed beetles a diet high in carbohydrate, conditions unlikely to induce gluconeogenesis. The effects of nutritional status on gluconeogenesis from alanine are known in M. sexta (Thompson, 1995). Last instar larvae administered (3-13C-alanine) display selective 13C enrichment in [1,2,5,6-13C]trehalose consistent with de novo carbohydrate synthesis. Gluconeogenesis is only evident in larvae maintained on a carbohydrate-free diet or a diet low in sucrose, compared with the sucrose level in normal rearing diets. 13C enrichment of trehalose from acetyl CoA formed by pyruvate decarboxylation and tricarboxylic acid cycle metabolism is less than 10% in glucogenic larvae. The ratio of pyruvate carboxylation/ decarboxylation is approximately 4, while this ratio in non-glucogenic larvae maintained on high carbohydrate diets is <1. Pyruvate cycling is markedly
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decreased in glucogenic larvae, indicating that pyruvate kinase plays an important role in regulating gluconeogenesis (Thompson, 2000). In this regard, regulation in insects may be similar to that observed in mammalian liver during starvation, but not in the liver of fed animals (Groen et al., 1983). Despite active carbohydrate synthesis via gluconeogenesis in M. sexta, haemolymph trehalose concentration is maintained at low levels, often <10 mM, in the absence of dietary carbohydrate. Such low haemolymph trehalose concentrations indicate a need for dietary carbohydrate and influence dietary selection behaviour (see Section 7.3). Under these conditions, haemolymph trehalose level is positively correlated to dietary protein consumption (Thompson et al., 2003) (Fig. 4 right). Investigations with low carbohydrate diets demonstrate than larvae consuming less than approximately 75 mg/day of carbohydrate are gluconeogenic, with haemolymph trehalose levels increasing with increased sucrose consumption (Thompson et al., 2002, 2003). Gluconeogenesis is under long-term regulation (Thompson, 1998). When larvae are transferred from a high carbohydrate diet to a carbohydratefree diet, gluconeogenesis is induced within several hours, and is maximal after 48 h. Infusion of cordycepin, a transcription inhibitor, inhibits gluconeogenesis, suggesting that regulation through gene expression and enzyme synthesis is the principal mechanism of nutritional regulation. Interestingly, gluconeogenesis in M. sexta may lack short-term regulation in reponse to dietary sucrose or haemolymph glucose or trehalose levels. Gluconeogenic larvae maintained on a carbohydrate-free diet fail to down-regulate gluconeogenesis in response to short-term sucrose feeding or by injection of glucose, even though haemolymph trehalose level increases in response to either treatment (Thompson, 1997a). An absence of short-term regulation over trehalose synthesis may explain why haemolymph trehalose is so variable under certain dietary conditions. Activity of the enzymes of gluconeogenesis have been examined in fat body preparations of the American cockroach (Storey and Bailey, 1978a). The cytosolic activities of fructose-1,6-bisphosphatase and phosphoenolpyruvate carboxykinase are much higher than that of pyruvate carboxylase, which is found exclusively in the mitochondrion. Glucose-6-phosphatase is associated with the microsomal fraction but its activity is relatively lower, and similar to that of pyruvate carboxylase. Pyruvate carboxylase and fructose-1,6-bisphosphatase have been purified from flight muscle of several insects, but the muscle enzymes have different properties from those of the fat body (Surholt et al., 1990; Tu and Hagedron, 1992). Flight muscle is a non-gluconeogenic tissue, and pyruvate carboxylase appears to function in supplying oxaloacetate for tricarboxylic acid cycle activation during flight initiation. Fructose-1,6-bisphosphatase has long been considered to provide for non-shivering thermogenesis through increased fructose phosphate cycling prior to flight. This role, however, has recently been disputed (Heinrich, 1993).
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4.1.4.2 Gluconeogenesis from glycerol. Glycogen formation by gluconeogenesis from dietary glycerol occurs in starved female A. taeniorhynchus mosquitoes (Nayar and Sauerman, 1971). Glycogen and trehalose are synthesized from tritiated glycerol fed to Pieris brassicase cabbage butterfly larvae (Turunen, 1993). In M. sexta larvae maintained on low carbohydrate diets, trehalose synthesis from (1,3-13C2)glycerol occurs at rates estimated between 0.5 and 3 mol glucose/g fresh weight, similar to those reported by others for fasting man (Thompson, 1997b). A special case of fat body glycogen synthesis via gluconeogenesis from 14Cenriched glycerol occurs in diapausing Chilo suppressalis larvae maintained at 25 C, after larvae had accumulated glycerol in response to cold exposure (Tsumuki and Kanehisa, 1981). A similar conversion of glycerol to glycogen occurs in cold-hardy larvae of two other lepidopteran larvae, Epiblema scudderiana and Eurostra solidaginis, when larvae with high levels of glycerol, accumulated in response to cold exposure, are exposed to increasing temperature (Joanisse and Storey, 1994a; Holden and Storey, 1995) (see Section 4.2.2.1). The latter authors reported that catabolism of glycerol to triose phosphate by these insects involves the activities of polyol dehydrogenase and glyceraldehyde kinase, rather than glycerol kinase. Glyceraldehyde kinase is a novel enzyme. 4.1.5
Biosynthesis from glycogen
Trehalose formation from fat body glycogen is well documented. Glycogen breakdown during exercise or starvation is mediated by a hormonally induced enzyme cascade similar to that known in vertebrate liver, where ultimately inactive glycogen phosphorylase b is phosphorylated to active glycogen phosphorylase a (EC 2.4.1.1) (see Sections 4.1.6 and 5.1.1) (Fig. 9). The properties of insect glycogen phosphorylases were reviewed by Steele (1982). Muscle phosphorylase plays little direct role in affecting haemolymph trehalose level, as it is active during periods of muscular activity when the glucose produced is immediately consumed for production of ATP. Activation and/or deactivation of the fat body enzyme has been described in the migratory locust and American cockroach, as well as in larvae or pupae of several lepidopteran species, Samia cynthia, B. mori, H. cecropia and recently M. sexta. Glycogen is rapidly hydrolysed during starvation of lepidopteran larvae. In B. mori, active fat body glycogen phosphorylase activity increases significantly within a few hours starvation, and haemolymph trehalose concentration increases for several hours as fat body and midgut glycogen content gradually decrease (Satake et al., 2000) (Table 2). Glycogen degradation has also been examined in M. sexta larvae starved for several days (Siegert, 1987; Gies et al., 1988). In those longer-term experiments, increased activity of glycogen phosphorylase correlates with decreased fat body glycogen. Haemolymph trehalose level increases slightly for several hours, remains within the normal
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range for fed animals, 25–40 mM, for at least 48 h, and thereafter decreases (Table 2). Glycogen phosphorylase is inactivated after approximately 24 h of starvation (Siegert et al., 1982; Gies et al., 1988). In both B. mori and M. sexta, haemolymph glucose concentration decreases sharply from the onset of starvation. Absorption of dietary sugar undoubtedly contributes to the maintenance of trehalose during starvation of these species, as larvae were well fed beforehand. If newly moulted unfed larvae are starved, haemolymph trehalose concentration decreases precipitously (Dahlman, 1973; Thompson, 2000). Care is required in interpreting results of studies with starved insects. In some cases, insects were denied both food and water, while in others water was provided. Small but significant differences between insects given or denied water are in part due to changes in haemolymph volume (Woodring, 1984). Feeding sucrose to starved M. sexta larvae results in inactivation of glycogen phosphorylase (Siegert and Mordue, 1992). The inactivation occurs quickly before any significant increase in haemolymph glucose or trehalose levels is evident, suggesting that a factor released from the gut in response to feeding may play a role. A similar inactivation of phosphorylase occurs if glucose is injected into larvae, but trehalose is ineffective. This suggests that regulation of glycogen phosphorylase may involve glucose sensitive control by the brain. Starvation or flight of adult M. sexta also results in fat body glycogen breakdown and trehalose formation (Ziegler and Schulz, 1986; Ziegler, 1991). In this case, however, activation of glycogen phosphorylase does not appear to be under hormonal control, at least not by the same hormone as in larvae (see Section 5.1.1). 4.1.6
Role of the fructose phosphate cycle in facilitating biosynthesis from glycogen and via gluconeogenesis
The fructose phosphate substrate cycle is a principal step in regulating glycolysis and gluconeogenesis (van Schaftingen, 1990) (see Section 4.1.4.1). The reactions have important ramifications for trehalose synthesis because trehalose synthesis and glycolysis compete for the same substrates. Fructose2,6-bisphosphate is the principal effector, activating 6-phosphofructo-1-kinase and inhibiting fructose-1,6-bisphosphatase (Berg et al., 2002). The level of fructose-2,6-bisphosphate is regulated by the activities of two enzymes, 6phosphofructo-2-kinase and fructose-2,6-bisphosphatase. In mammals, both enzymes are present in a single bifunctional protein (Pilkis and Claus, 1995). The relationships between the synthesis and degradation of fructose-2,6bisphosphatase and the activities of 6-phosphofructo-1-kinase and fructose1,6-bisphosphatase in affecting flux through the fructose phosphate substrate cycle are illustrated in Fig. 7. The importance of fructose-2,6-bisphosphate in regulating carbohydrate metabolism in insects was first demonstrated in the cabbage looper Trichoplusia ni (Thompson, 1985). Fructose-2,6-bisphosphate, as well as AMP, inhibit
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FIG. 7 Metabolic scheme illustrating the regulation of the fructose phosphate substrate cycle by fructose-2,6-bisphosphate. The bold arrows indicate the glycolytic reaction catalysed by 6-phosphofructo-1-kinase (left), and the gluconeogenic reaction catalysed by fructose-1,6-bisphosphatase (right).
fructose-1,6-bisphosphatase activity in the soluble fraction of fat body homogenates. Moreover, inhibition by each effector is enhanced in the presence of the other, a finding consistent with the synergistic role of fructose-2,6-bisphosphate in mediating the effect of AMP in liver (van Schaftingen and Hers, 1981). Due to a decrease in the sensitivity of fructose-1,6-bisphosphatase to both fructose-2,6bisphosphate and AMP, larvae parasitized by the ichneumonid Hyposoter exigua display increased gluconeogenesis and an elevated haemolymph trehalose level (Thompson and Binder, 1984). Fructose-2,6-bisphosphate also regulates trehalose synthesis in the Argentine cockroach, Blaptica dubia. In response to hormonally mediated decreases in fructose-2,6-bisphosphate, synthesis of trehalose by incubated fat body increases (Becker and Wegener, 1998) (see Section 5.1.1). The effect is accompanied by a significant decrease in 6-phosphofructo-1-kinase activity and a simultaneous increase in fructose-1,6-bisphosphatase activity (Becker et al., 1996) (Fig. 8). Glycogen was assumed to provide the glucose for trehalose synthesis, but gluconeogenesis could be involved as well. In the cold-hardy lepidopterans E. solidaginis and E. scudderiana, the enzymes of the fructose phosphate cycle regulate the synthesis of trehalose
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FIG. 8 Inverse effect of fructose-2,6-bisphosphate on 6-phosphofructo-1-kinase and fructose-1,6-bisphosphate activities in fat body of Blaptica dubia. (Data after Becker et al., 1996) Assay conditions as outlined in that report.
and the reformation of glycogen from glycerol via gluconeogenesis (Holden and Storey, 1993, 1995). Activity of 6-phosphofructo-1-kinase in E. solidaginis is inhibited by increasing temperature, while fructose-1,6-bisphosphatase activity is stimulated. Accompanying the temperature-dependent activation of fructose-1,6-bisphosphatase are significant increases in the KI (inhibitor concentration for approximately half-maximal velocity) for fructose-2,6bisphosphate and AMP. Between 5 and 22 C, values for these effectors increase 8 fold and 2.5 fold, respectively. Moreover, the inhibition by fructose2,6-bisphosphate is mediated by glycerol, with no inhibition evident at 22 C in the presence of 2 M glycerol. Subsequent studies with fructose-1,6bisphosphatase purified from E. scudderiana, demonstrates that activity is in part dependent on the reversible phosphorylation of the enzyme, with the phosphorylated enzyme the principal form present under warm temperature conditions favouring gluconeogenesis (Muise and Storey, 1997). The phosphorylated form exhibits much lower sensitivity to fructose-2,6-bisphosphate and AMP inhibition than the non-phosphorylated enzyme, which is present in a greater proportion in cold conditions. The hormonal regulation of the fructose phosphate cycle is described below (see Section 5.1.1). 4.2 4.2.1
DEGRADATION
Hydrolysis
Trehalase (EC 3.2.1.28) catalyses the irreversible hydrolysis of trehalose to glucose. As the only known enzyme for metabolizing trehalose to glucose, it is not surprising that trehalase activity is found in many forms and is widely
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distributed among insect tissues and organs, including haemolymph. Numerous membrane-bound and soluble isozymes have been isolated, including enzymes from H. cecropia (Gussin and Wyatt, 1965) P. regina (Hansen, 1966), B. mori (Saito, 1960; Egorova and Khomidov, 1991), P. americana (Takahashi et al., 1980; Jahagirdar et al., 1990), A. mellifera (Talbot and Huber, 1975; Lee et al., 2001) as well as many other insects reviewed previously (Friedman, 1985). The reaction mechanism and stereochemistry were described by Clifford (1980) and Labat-Robert et al. (1978), respectively. The kinetics and conditions for optimal activity vary greatly between enzymes from different insects and tissues. Trehalase genes have been characterized from B. mori (Su et al., 1993, 1994) and T. molitor (Takiguchi et al., 1992). Baculovirus-mediated expression of the T. molitor gene has been achieved in the cabbage armyworm Mamestra brassicae and in an insect cell line, SF-9, derived from Spodoptera frugiperda, the fall armyworm (Sato et al., 1997). Trehalose hydrolysis is perhaps best understood in asynchronous flight muscle of dipteran insects, many of which utilize trehalose as the principal source of energy during flight. In P. regina, the enzyme is localized in the inner membrane of the mitochondria (Reed and Sacktor, 1971), and there is a strong correlation between wingbeat frequency and decreased haemolymph trehalose level (Clegg and Evans, 1961). Sacktor and Wormser-Shavit (1966) demonstrated that trehalose concentration in the thorax decreases sharply during the first few seconds of flight, and thereafter decreases continuously but at a lesser rate. Glucose increased for approximately 30 s before decreasing due to oxidation. Trehalase in flight muscle homogenate displays an activity of 0.2 mol glucose formed/min/mg protein (Reed and Sacktor, 1971), a level sufficient to account for the rate of trehalose utilization. Insects with synchronous flight muscle, such as H. cecropia (Gussin and Wyatt, 1965) the cockroach, Blaberus discoidalis (Gilby et al., 1967) and the desert and migratory locusts (Vaandrager et al., 1989), utilize trehalose and fat as fuels for flight. Trehalase in these insects is membrane-bound and associated with the microsomal cell fraction. Studies with American cockroach confirm the critical role of trehalase in trehalose hydrolysis during flight (Kono et al., 1994). When validoxylamine, an amino sugar and potent competitive inhibitor of trehalase, is injected into adult males, trehalase activity is dramatically reduced and trehalose accumulates approximately 3-fold in the haemolymph. Very active insects are able to fly only a few minutes after treatment, compared with 15 min for untreated individuals. Subsequent studies with three dipteran species, Musca domestica, Boettcherisca peregrina and Calliphora nigribarbis, also demonstrate inhibition of trehalase and accumulation of trehalose following validoxylamine treatment (Takahashi et al., 1995). The flight activity of M. domestica and B. peregrina is reduced by validoxylamine treatment, but flight activity in the blow fly C. nigribarbis is unaffected or increased. The authors concluded that the latter species is able to use an alternate fuel source, possibly lipid, for flight. Because other investigations demonstrate that
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C. nigribarbis displays long distance flight and migration (Kurahashi et al., 1991), it may be that this species has an unusual pattern of fuel utilization. As described above, many dipterans have significant levels of haemolymph proline that is utilized as fuel. Although proline is generally of lesser importance than glucose as a fuel in the blow fly, proline or other amino acids may provide fuel if glucose is unavailable. Trehalase activity is present at relatively high levels in insect midgut. There, it may aid in maintaining the glucose concentration gradient across the midgut epithelium, by hydrolysing trehalose, and preventing the diffusion of trehalose from the haemolymph into the gut lumen (Wyatt, 1967). Trehalase serves as a digestive enzyme in insects feeding on foods containing trehalose (Terra and Ferreira, 1994). The enzyme in B. mori larval midgut is associated with the basal plasma membrane and not the microvilli, lending support to the former role in this species (Azuma and Yamashita, 1985). The extracellular surfaces of the follicle cells of B. mori ovaries producing diapausing eggs have a high level of trehalase activity (Shimada and Yamashita, 1979). This is associated with trehalose hydrolysis and the provision of glucose for the synthesis and accumulation of glycogen in ovarian cells (Yamashita et al., 1972). The process is under hormonal control (see Section 5.4). Inhibition of trehalase with the inhibitor trehazolin prevents glycogen accumulation but does not affect embryogenesis (Katagiri et al., 1998). In contrast, oocyte development in P. americana is suppressed if trehalase is inhibited with validoxylamine (Kono et al., 1997). An interesting question arises due to the occurrence of trehalase in haemolymph (van Handel, 1978). Because trehalase and trehalose are both present, how are trehalase activity and trehalose hydrolysis regulated? Compartmentation may be one mechanism, if trehalase is released from haemocytes. Studies with American cockroach demonstrate that pH plays a role, as trehalase is activated by decreases in haemolymph pH associated with physical activity (Downer and Matthews, 1977). Diet and starvation can have dramatic effects on haemolymph pH (Harrison, 2001) and may, therefore, be involved in regulating trehalase. Hormones or other regulatory factors likely play an important role as well. A haemolymph peptide that inhibits trehalase has been isolated and characterized from American cockroach (Hayakawa et al., 1989), and studies with numerous insect species demonstrate the presence of hypotrehalosemic insulin-like peptides that activate trehalase (see Section 5.2). 4.2.2
Catabolism
4.2.2.1 Glycolysis and formation of polyols. Insects catabolize glucose derived from trehalose or glycogen via the well-known Embden–Meyerhof glycolytic pathway or the pentose phosphate pathway discussed above (see Sections 4.1.3 and 4.1.4.1). Glycolysis has been examined in a variety of
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insect tissues, and is especially well documented in flight muscle. The activities and regulatory properties of the enzymes were reviewed previously (Friedman, 1985). Collectively, these investigations demonstrate a high glycolytic flux and metabolic rate in insects utilizing carbohydrate as the principal fuel for flight. The metabolic rate is closely correlated with the activities of hexokinase and 6-phosphofructo-1-kinase, the two principal regulatory enzymes of the glycolytic pathway (Crabtree and Newsholme, 1975; Sacktor, 1975; Steele, 1981; Suarez, 2000). During flight of the honeybee, these two enzymes operate at 76% and 44%, respectively, of their maximal capacity (Vmax) (Suarez et al., 1996). Glycolytic enzymes from fat body have also been examined, and often have different properties from the muscle enzymes. Three isozymes of hexokinase, for instance, occur in fat body of B. mori larvae, each having different affinities for ATP and glucose, compared with only one form in muscle (Yanagawa, 1978). On the other hand, fat body 6-phosphofructo-1-kinase from S. gregaria is similar to the muscle enzyme although the affinity for fructose-6-phosphate appears to be lower in fat body (Walker and Bailey, 1969). Both the fat body and the muscle enzymes are inhibited by ATP and stimulated by AMP, and both enzymes are inhibited by fructose-1,6-bisphosphate, an activator of the enzyme in mammalian muscle. Moreover, while cAMP stimulates the muscle enzyme of the desert locust, the fat body enzyme is unaffected. As noted previously (Friedman, 1985), the activity of several glycolytic enzymes, including 6-phosphofructo-1-kinase, in fat body of American cockroach are very low relative to that of glucose-6-phosphate dehydrogenase (Storey and Bailey, 1978a,b), which catalyses the first step of the pentose phosphate pathway (Fig. 5). This further suggests that the pentose phosphate pathway plays a principal role in glucose oxidation by fat body (see Section 4.1.3). The fat body enzymes of the fructose phosphate substrate cycle have been examined in two species, T. ni (Thompson and Binder, 1984) and B. dubia (Becker et al. 2001). The kinetic behaviour of 6-phosphofructo-1-kinase is similar in both species, and stimulated by AMP. The effect is mediated by ATP for the T. ni enzyme, and in the absence of AMP, ATP is inhibitory above 0.5 mM. 6-phosphofructo-1-kinase from B. dubia is activated by fructose2,6-bisphosphate, as described above (see Section 4.1.6), and as is the case with the desert locust, fructose-1,6-bisphosphate inhibits the activity of 6-phosphofructo-1-kinase in B. dubia. The effects of fructose-2,6-bisphosphate and fructose-1,6-bisphosphate were not examined for the enzyme from T. ni. The kinetic behaviour of fructose-1,6-bisphosphatase in response to fructose-1,6-bisphosphate is similar for the enzymes from both T. ni and B. dubia, with this substrate inhibitory above approximately 50 mM. The enzyme from both species is inhibited by fructose-2,6-bisphosphate and AMP. Fat body pyruvate kinase has been characterized from S. gregaria (Bailey and Walker, 1969). The enzyme is activated by fructose-1,6-bisphosphate at concentrations much lower than with the enzyme derived from muscle, but
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the enzymes from both tissues are inhibited by ADP above 10 mM, as well as by ATP. Inhibition of the fat body enzyme by ADP and ATP is relieved by fructose-1,6-bisphosphate. The maximal activities of all the glycolytic enzymes of fat body of P. americana were examined by Storey and Bailey (1978b). The activity of pyruvate kinase greatly exceeds that of the other enzymes catalyzing the rate-limiting steps of glycolysis, including glycogen phosphorylase, hexokinase and 6-phosphofructo-1-kinase. The potential role of pyruvate kinase in regulating gluconeogenesis in M. sexta is described above (see Section 4.1.4.1). The synthesis of glycerol and various sugar alcohols including sorbitol, ribitol, erythritol, threitol and mannitol from glycogen and/or trehalose is well documented in insects (Storey and Storey, 1991; Storey, 1997). Accumulation of these metabolites is most often associated with cold-hardiness, survival under low temperature due to freeze tolerance or the avoidance of freezing in response to cold exposure (Storey and Storey, 1988; Lee, 1991) (see Section 7.2.1). Cold-hardiness is sometimes associated with diapause (Denlinger, 1991). Most cold-hardy insects synthesize polyols solely from glycogen. Trehalose is not involved and often remains at a relatively constant level in the haemolymph. This metabolism, first described in diapausing embryos of B. mori eggs (Chino, 1961, 1963), is now known in numerous insects (Lee, 1991). Glycerol synthesis in B. mori and many other species involves the glycolytic reactions leading to the triose phosphates. Dihydroxyacetone phosphate is reduced to -glycerophosphate catalysed by cytoplasmic -glycerophosphate dehydrogenase (EC 1.1.1.8) and alkaline phosphatase (EC 3.1.3.1)-catalysed dephosphorylation to glycerol (Yaginuma and Yamashita, 1980) (Fig. 5). Polyol accumulation is usually accompanied by a decrease in mitochondrial activity and aerobic respiration (Joanisse and Storey, 1994b). Cold-induced mitochondrial degradation occurs during glycerol accumulation in some insects (Kukal et al., 1989). The -glycerophosphate dehydrogenase-catalysed reduction of dihydroxyacetone phosphate simultaneously reoxidizes NADH þ H þ to NAD þ , thereby maintaining redox balance in the cytosol. This same cytosolic enzyme carries out the cytoplasmic portion of the shuttle transferring reducing equivalents into the mitochondrion during aerobic respiration. Some variation in the fate of the triose phosphates is apparent between species. In larvae of E. solidaginis, glycerol may arise from glyceraldehyde-3phosphate through the actions of glyceraldehyde phosphatase and NADP þ dependent polyol dehydrogenase (Joanisse and Storey, 1994a). Sorbitol is synthesized directly from glucose in an aldose reductase (EC 1.1.1.21)-catalysed reaction that regenerates NADP þ (Fig. 5). Alternately, sorbitol may arise from glucose-6-phosphate through action of a polyol phosphate dehydrogenasecatalysed reaction that also regenerates NADP þ , followed by dephosphorylation to sorbitol, catalysed by alkaline phosphatase (EC 3.1.3.1). Oxygen availability influences the pattern of polyol synthesis. Glycerol synthesis requires ATP during the formation of fructose-1,6-bisphosphate,
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while sorbitol production does not require energy. Thus, for instance in B. mori eggs, glycerol is synthesized when diapause is initiated and oxygen is available. As the serosal cuticle is formed and oxygen becomes limiting, sorbitol is preferentially formed (Sonobe et al., 1979). Regulation of polyol formation occurs at several steps. Hayakawa et al., (1989) reported a significantly reduced activity of glycogen phosphorylase phosphatase, the enzyme catalysing the deactivation of glycogen phosphorylase, in fat body of Philosamia cynthia silkworm pupae exposed to cold. Pfister and Storey (2002) described the behaviour and inhibition by cold of the same enzyme, now known as protein phosphatase-1, purified from larvae of E. scudderiana and E. solidaginis. The cold-induced activation of glycogen phosphorylase in these species is accompanied by a simultaneous inhibition of glycogen synthase (Hawakawa and Chino, 1982a; Muise and Storey, 1999). Indeed, polyol synthesis during cold exposure, and reformation of glycogen from polyols when warmer temperatures prevail, are principally due to the reciprocal activation and deactivation of glycogen phosphorylase and glycogen synthase (Fig. 9). Dramatic changes in regulatory properties in response to temperature have also been reported for partially purified hexokinase isolated from E. scudderiana (Muise and Storey, 2001). Enzymes of the fructose phosphate cycle also appear to play a major role. 6-phosphofructo-1-kinase was implicated by Hawakawa and Chino (1982b). Activity of the enzyme increases several fold during diapause of Papilio machaon pupae and Monema flavescens prepupae, two lepidopterans that accumulate glycerol, compared with Philosamia cynthia pupae and Trichiocamps populi prepupae, species that accumulate trehalose. 6-phosphofructo-1-kinase purified from E. scudderiana is cold activated, thereby facilitating the conversion of glycogen to triose phosphates and subsequently to glycerol (Holden and Storey, 1993). Stimulation of enzyme activity by the effectors fructose-2,6-bisphospate and AMP is strongly temperature-dependent. Considerable evidence demonstrates that the pentose phosphate pathway is not only the source of several sugar alcohols, but also provides the reduced power for polyol formation during cold exposure. In the absence of the oxidative phase of glycolysis and of oxidation of fatty acids, the pentose phosphate pathway is the sole source for reduced coenzyme. The approach employed to examine the pentose phosphate pathway compared the oxidation of 14C-enriched isotopomers of glucose, as described above (Section 4.1.3). Estimating the contribution of the pentose phosphate pathway to glucose oxidation based on product yields is greatly simplified, because aerobic respiration is significantly reduced during cold exposure. Increased pathway activity during polyol synthesis occurs in diapausing B. mori eggs (Kageyama, 1976), the adult blow fly Protophormia terranovae (Wood and Nordin, 1980), and cold-hardy larvae of E. solidaginis (Tsumuki et al., 1987). The stoichiometry of glycerol and sorbitol production in relation to glycolytic and pentose phosphate pathway activity was outlined by Storey and
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FIG. 9 Metabolic scheme illustrating the reciprocal activation and deactivation of glycogen phosphorylase and glycogen synthase in insects. (Upper) Enzyme cascade activating glycogen phosphorylase and glycogen synthase and the role of effectors in regulation. (Lower) Interaction of glycogen phosphorylase and glycogen synthase in regulating polyol metabolism in cold-hardy insects. (After Storey and Storey, 1991.)
Storey (1983). The formation of 12 moles of glycerol from 6 moles of glucose-6phosphate requires 12 moles of NADPH þ H þ , necessitating the oxidation by the pentose phosphate pathway of the equivalent of 1 mole of glucose-6phosphate. In addition, 5 moles of ATP are required for the synthesis of 10 moles of triose phosphate, the equivalent of 2 moles of triose phosphate being provided by the pentose phosphate pathway. This energy requirement necessitates the oxidation of approximately 0.13 (5/38) moles glucose by aerobic respiration. The situation with sorbitol is simpler, as the synthesis of 1 mole sorbitol from glucose or glucose-6-phosphate requires 1 mole of NADPH þ H þ and ATP is not required. In this case, however, cycling to glucose-6-phosphate occurs. Overall efficiency of sorbitol synthesis was estimated at 92%, with 8% of carbon lost by oxidation in the pentose phosphate pathway. Efficiency for glycerol formation was 84%. In vivo studies demonstrated that the formation of glycerol and sorbitol by E. solidaginis larvae is 93% efficient at two temperatures, 0 and 30 C (Storey et al., 1981).
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In contrast to polyol synthesis, less is known concerning the reactions and enzymes that catalyse the reoxidation and metabolism of polyols to glycolytic intermediates and/or glucose and glycogen. The enzyme catalysing the oxidation of sorbitol to glucose, sorbitol dehydrogenase, is partially characterized from B. mori, and is NAD þ specific (Yaginuma and Yamashita, 1979). Sorbitol metabolism in E. solidaginis also involves sorbitol dehydrogenase, followed by the action of hexokinase to yield glucose (Joanisse and Storey, 1994a). The special case where glycogen is resynthesized from glycerol via gluconeogenesis in cold-hardy insects is described above (see Section 4.1.4.2). The activities of glycogen phosphorylase and glycogen synthase upon resynthesis of glycogen from polyols are summarized in Fig. 9. 4.2.2.2 Aerobic respiration. Carbohydrates and fats are the principal substrates providing energy through aerobic respiration. Insects that oxidize trehalose and glycogen for flight exhibit the highest mass-specific rates of aerobic respiration known (Sacktor, 1975, 1976; Candy, 1989; Suarez, 2000). Differences in metabolic rates between rest and flight may be two orders of magnitude, with oxygen consumption during flight orders of magnitude greater than oxygen consumption by mammals at maximal activity (Weis-Fogh, 1964; Ziegler, 1984). Moreover, at rest following flight, ‘respiration of recovery’ is absent, or nearly so. During and immediately following flight initiation, most insects oxidize trehalose as an energy source to sustain aerobic respiration while glycogen and/or lipid are mobilized (Candy et al., 1997).
5
Hormonal regulation of metabolism
5.1 5.1.1
HYPERTREHALOSEMIC FACTORS
Hypertrehalosemic hormone and glycogen degradation
Steele (1963) was the first to demonstrate that extracts of the corpora cardiaca injected into the American cockroach stimulate glycogen breakdown and release of trehalose from the fat body into the haemolymph. Those findings were soon confirmed with a second cockroach, B. discoidalis (Bowers and Friedman, 1963). By comparison with glucagon, Steele (1980) suggested the name trehalogon for the factors responsible, but they are generally known as hypertrehalosemic hormones. Hypertrehalosemic hormones have now been demonstrated or isolated from many insects (Ga¨de, 1990, 1991; Ga¨de et al., 1994; Nijhout, 1994). They are octa-, nona- and decapeptides with several common structural features necessary for activity (Ga¨de, 1990; Ziegler et al., 1991). The hormones do not always produce an elevation in haemolymph trehalose level, but those that affect carbohydrate metabolism all act by stimulating glycogen breakdown (Goldsworthy and Ga¨de, 1983). In some
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insects, for example locusts, the same hormone acts to stimulate lipolysis, the hydrolysis of triglycerides, in addition to glycogen breakdown. Thus, the peptides are also known as adipokinetic hormones (Orchard, 1987; Ga¨de, 1990). Some insect species have more than one hormone. Each may exhibit different behaviour affecting different metabolism, or both may display similar activity. In M. sexta, the same peptide acts as hypertrehalosemic hormone in the larval stages and adipokinetic hormone in the adult (Ziegler and Schulz, 1986; Ziegler et al., 1990; Arrese et al., 1996). By convention, specific peptides are often referred to by specific names, hypertrehalosemic hormone or adipokinetic hormone, despite the fact that they may affect both lipid and carbohydrate metabolism, or affect the same metabolism in different species. The adipokinetic hormone of B. mori larvae is identical to the hypertrehalosemic hormone of M. sexta (Oda et al., 2000). The hypertrehalosemic and adipokinetic hormones of locusts and cockroaches are released from the corpora cardiaca upon initiation of and during exercise and flight (Orchard, 1987). They act to provide substrate, glucose and/or fatty acids, for supporting aerobic respiration. Hypertrehalosemic hormones stimulate glycogen hydrolysis in a similar manner as the mammalian hormones glucagon and epinephrine, as well as another insect hormone, octopamine. They act on the protein kinase enzyme cascade that activates phosphorylase kinase, and ultimately glycogen phosphorylase (Sutherland, 1972; Berg et al., 2002) (Fig. 9). The activities of hypertrehalosemic and adipokinetic hormones are mediated through several signal transduction pathways. In locusts, hormone-stimulated glycogenolysis involves both a cAMP/G protein-coupled pathway (Wang et al., 1990; Vroemen et al., 1995), as well as extracellular and intracellular Ca2 þ (van Marrewijk et al., 1993). Hypertrehalosemic action in cockroaches, however, does not involve cyclic nucleotides (Ga¨de et al., 1994). These differences between species occur despite the structural similarities of the hormones involved. The signal transduction pathway in cockroaches involves inositol triphosphate stimulated release of intracellular Ca2 þ , and influx of extracellular Ca2 þ for maximal response (Steele and Paul, 1985; Lee and Keeley, 1994; Keeley et al., 1996; Park and Keeley, 1996a; Steele et al., 2001; Sun et al., 2002). Clearly, the actions by different adipokinetic and hypertrehalosemic hormones in different species involve multiple receptors and signal transduction pathways, as is the case with (nor)epinephrine, vasopressin and other mammalian hormones (Bolander, 1994). The hypertrehalosemic hormone of B. discoidalis simultaneously stimulates glycogen phosphorylase and trehalose-6-phosphate synthase (Keeley et al., 1996). In cockroaches, activation of glycogen phosphorylase appears to be the rate-limiting step in trehalose synthesis from glycogen (Steele and Hall, 1985). Studies with B. discoidalis suggest that the actions of hypertrehalosemic hormones on trehalose synthesis may be coordinated by eicosanoids (Keeley et al., 1996). Addition of an inhibitor of arachidonic acid metabolism enhances the
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effect of hypertrehalosemic hormone on trehalose synthesis in cockroach fat body preparations. Studies with P. americana also indicate a role for arachidonic acid metabolites in mediating the actions of hypertrehalosemic hormone (Ali et al., 1998). Cyclooxygenase inhibitors strongly inhibit the activation of trehalose efflux by hypertrehalosemic hormone. This suggests that prostaglandins may be important in regulating trehalose synthesis and efflux, a conclusion supported by an earlier demonstration that prostaglandin F2 stimulates trehalose efflux from the fat body (Ali and Steele, 1997). The action of phospholipase A2 may generate the arachidonic acid required for prostaglandin synthesis (Ali et al., 1998). Little is known of the importance of hypertrehalosemic hormones in regulating other aspects of intermediary carbohydrate metabolism. In the Argentine cockroach, B. dubia, hypertrehalosemic hormone is clearly involved in regulating flux at the fructose phosphate substrate cycle (see Section 4.1.6). When fat body is incubated with extracts of corpora cardiaca containing the hormone, the level of the glycolytic activator, fructose-2,6-bisphosphate is decreased, and trehalose release into the medium is significantly increased (Becker and Wegener, 1998). Subsequent studies demonstrate that the decrease in fructose-2,6-bisphosphate simultaneously increases fat body fructose-1,6bisphosphatase and depresses 6-phosphofructo-1-kinase activities, and these effects are mediated through signal transduction pathways involving Ca2 þ (Becker et al., 1998, 2001). Extracts of the corpora cardiaca affect pentose phosphate pathway activity in L. migratoria (Gourdoux et al., 1985), T. molitor (Gourdoux, 1980; Mtioui et al., 1993a, 1994) and Pieris brassicae (Moreau et al., 1980; Gourdoux et al., 1989), reducing the contribution of this pathway to glucose oxidation relative to oxidation by glycolysis. Hypertrehalosemic hormone has not been shown to affect gluconeogenesis, although its effect on increasing fructose-1,6-bisphosphatase activity in the fat body of Argentine cockroach clearly suggests that this may be so. Injection of corpora cardiaca extract into American cockroach fails to induce gluconeogenesis and trehalose or glycogen synthesis from alanine (Sevala and Steele, 1989). In the latter investigation, however, insects were fed a sucrose solution before the experiments. Studies with B. discoidalis cockroaches demonstrate that starvation significantly enhances expression of hypertrehalosemic hormone mRNA transcript (Lewis et al., 1998). Synthesis and secretion of hypertrehalosemic hormone are also increased by starvation (Sowa et al., 1996). If hypertrehalosemic hormone induces gluconeogenesis in starved cockroaches, in addition to or following its effect on increasing trehalose formation from glycogen, the hormone is similar in activity to glucagon. On the other hand, hypertrehalosemic hormones that fail to affect gluconeogenesis would appear to be more epinephrine-like. Epinephrine does not affect gluconeogenesis directly. In this regard, future study may
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demonstrate differences between hypertrehalosemic and adipokinetic peptides among species. Substances in extracts of corpora cardiaca that are immunoreactive with polyclonal antibodies of known adipokinetic hormones are found in numerous insect species representing 10 insect orders (Schooneveld et al., 1987). This suggests that these homones are widely distributed among insects. The findings, however, must be interpreted with caution, as immunological results do not confirm the chemical structure of the peptides, but rather identify common epitopes present in the antibodies (Ga¨de, 1990). 5.1.2
Other hypertrehalosemic factors
The biogenic amine octopamine, an analogue of norepinephrine, is widely distributed in the nervous system of insects (Orchard et al., 1993; Roeder, 1999). When released into the haemolymph, octopamine acts as a neurohormone, having effects on carbohydrate and lipid metabolism. When excited, cockroaches such as P. americana exhibit short-term hypertrehalosemia in response to octopamine (Downer, 1979; Davenport and Evans, 1984). The effect is distinguished from that of hypertrehalosemic hormone by head ligation which eliminates action by the latter hormone (Downer, 1980; Woodring et al., 1989). Octopamine activates fat body glycogen phosphorylase in B. discoidalis (Park and Keeley, 1996b). Trehalase activity in muscle and haemolymph is increased by injection of octopamine into American cockroach (Jahagirdar et al., 1984). In the migratory locust, octopamine enhances trehalose utilization and glucose oxidation by flight muscle (Candy, 1978; Whim and Evans, 1988), and stimulates triglyceride hydrolysis leading to hyperlipidemia (Orchard et al., 1981). How responses to octopamine are coordinated with related responses due to the action of hypertrehalosemic and adipokinetic hormones in different insect species is complex and not well understood (Woodring et al., 1989; Orchard et al., 1993). Investigations with the migratory locust indicate that octopamine simultaneously acts as a neurotransmitter, regulating the release of adipokinetic hormone, as well as a neurohormone, stimulating fat body lipid hydrolysis. Octopamine injected into migratory locusts increases fructose-2,6-bisphosphatase, thereby increasing the activity of 6-phosphofructo-1-kinase and glycolysis (Blau and Wegener, 1994) (see Section 4.1.6). Similar effects are observed in isolated flight muscle perfused with octopamine (Blau et al., 1994). Adipokinetic hormones, on the other hand, have no effect on the level of fructose-2,6-bisphosphatase. Recent studies demonstrate that prior to flight octopamine released from modulatory dorsal unpaired median neurones innervating flight muscle acts to increase fructose-2,6-bisphosphate and activate glycolysis (Mentel et al., 2003). Following flight initiation, octopamine release declines, thereby reducing glycolysis and carbohydrate oxidation. Muscle and haemolymph octopamine levels appear compartmentalized, as
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haemolymph octopamine remains high while muscle levels decline (Orchard et al., 1993; Rheuben, 1995). Presumably, at this point octopamine acts in concert with adipokinetic hormone to stimulate lipid oxidation for prolonged flight. Like hypertrehalosemic and adipokinetic hormones, octopamine actions involve multiple signal transduction pathways and receptor types (Nathanson, 1993; Mentel et al., 2003). Extracts of various structures from insect brain or nervous system contain compounds that are immunologically reactive to glucagon antibodies (Kramer, 1980, 1985). Indeed, hypertrehalosemic hormone is such an immunoreactive glucagon-like peptide, despite the size and structural dissimilarity of glucagon and hypertrehalosemic hormone, and the fact that glucagon generally fails to elicit similar responses in insects as hypertrehalosemic hormones. Nevertheless, because of the large number of such glucagon-like compounds, the immunological results suggest that insects may have additional hormones acting on carbohydrate and trehalose metabolism. Numerous such compounds are found circulating in the haemolymph, but their functions are generally unknown (Kramer et al., 1980). Studies with M. sexta demonstrate the presence of a high molecular weight glucagon-like peptide in the midgut (Tager and Kramer, 1980). Indeed, insects are now known to have a complex gut endocrine system, but little in known of the possible functions of the numerous peptides present (Sehnal and Zitnan, 1990) (see Section 7.3). 5.2
HYPOTREHALOSEMIC FACTORS
The occurrence in insects of compounds with hypotrehalosemic activity was first suggested from studies of the blow fly, C. erythrocephala, where decapitation results in a decrease in haemolymph trehalose concentration (Normann, 1975). The median neurosecretory cells of the brain appear to be the source of the factor responsible (Duve, 1978). Removal of the median neurosecretory cells from the brain causes a rise in the haemolymph concentrations of both trehalose and glucose, and when an extract of the cells is injected, the levels of both sugars decrease below normal (Table 2). Injection of the extract into intact flies causes a decrease in glucose, but trehalose concentration increases slightly. Similar results with P. regina were reported by Chen and Friedman (1977a,b), who demonstrated that removal of the corpora cardiaca/corpora allata produces an elevation of haemolymph trehalose, and that the effect is not due to glycogen breakdown. Further, trehalose synthesis is impaired in insects lacking corpora cardiaca/corpora allata, suggesting an insulin-like effect of reducing glucose uptake by the fat body. Trehalose accumulates in the haemolymph because of feedback inhibition of trehalase by glucose. Immunological investigations of insects from several orders, including Coleoptera, Diptera, Hymenoptera, Lepidoptera and Orthoptera, demonstrate
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the presence of insulin-like peptides and proteins in a variety of tissues (Kramer, 1985; Sevala et al., 1993). This suggests that related molecules are likely present in most insects. The insulin-like protein from haemolymph of M. sexta displays a close similarity in amino acid composition with porcine insulin (Kramer et al., 1982). Many insulin-like molecules in insect brain are immunologically associated with the prothoracicotropic hormones, hormones synthesized in medial neurosecretory cells involved in regulating activity of the prothoracic glands (Nijhout, 1994). Bombyxins, a family of at least three insulin-like peptides from B. mori, are sometimes referred to as ‘small’ prothoracicotropic hormone, although they actually exhibit limited prothoracicotropic activity in B. mori (Nagasawa, et al., 1990; Iwami, 2000). Insulin-like molecules, however, do regulate specific aspects of development in some species (Sevala and Loughton, 1992; Sevala et al., 1993; Leevers, 2001). Bombyxins share close sequence homology (Nagata et al., 1995a; Nagasawa et al., 1986), but are not structurally identical (Nagata et al., 1995a) to vertebrate insulin. They are known to be produced in several other insect tissues in addition to brain (Iwami et al., 1996). Extensive studies have been made on the expression of the bombyxin gene (Iwami, 1990; Iwami et al., 1996; Salam et al., 2001), and the bombyxin receptor on the surface of the prothoracic glands has been characterized (Nagata et al., 1995b). In the fruit fly Drosophila melanogaster, at least seven genes express insulinlike proteins in salivary gland, midgut, neurones of the ventral nerve cord and neurosecretory cells in the brain (Brogiolo et al., 2001; Cao and Brown, 2001; Rulifson et al., 2002). Gene expression occurs in a tissue- and stage-specific manner to regulate growth and development. In the larval stage, gene expression occurs predominantly in bilateral symmetric clusters of neurosecretrory cells located in the pars intercerebralis region of the brain (Rulifson et al., 2002). Processes of these cells extend to the corpora cardiaca of the ring gland as well as to the heart. Further, adipokinetic hormone expressing cells of the corpora cardiaca contain insulin-like peptide. Receptors for insulin-like proteins in fruit fly are characterized (Brogiolo et al., 2001; Kulansky Poltilove et al., 2001). Depending on the source and the molecule, insulin-like proteins act on carbohydrate and/or lipid metabolism in insects, as does vertebrate insulin on insects (Kramer, 1985). Bombyxin activates trehalase and reduces both haemolymph trehalose and fat body glycogen levels in B. mori larvae, although not in adults (Satake et al., 1997, 1999). Release of bombyxin from the brain of larvae is reduced by starvation, and restored upon refeeding (Masumura et al., 2000). Increased bombyxin release is also observed after injection of glucose into starved larvae, indicating that glucose may function as a regulator of haemolymph hormone level. An insulin-like peptide isolated from insect gut has a significant effect on glucose oxidation by the migratory locust (Ben Khay et al., 1987). The peptide, which is immunoreactive with vertebrate insulin antibodies, increases the
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activity of the pentose phosphate pathway relative to glycolysis when injected into adult locusts. The effect is mimicked by mammalian insulin, and is opposite that observed with locusts treated with extracts of corpora cardiaca containing hypertrehalosemic activity (Gourdoux et al., 1985). Similar effects of gut insulin-like peptide on pentose pathway activity occur in isolated fat body and muscle of T. molitor larvae (Mtioui et al., 1994). The effect of the hormone is mediated by cAMP signal transduction that involves influx of extracellular Ca2 þ (Mtioui et al., 1993a, 1996). Fat body trehalase is also activated by insulin-like peptide in T. molitor (Bounias et al., 1993). Ablation of the insulin-like protein secreting cells in the brain of D. melanogaster produces a significant elevation in the combined level of trehalose and glucose (Rulifson et al., 2002). Unfortunately, the two sugars were not distinguished during this investigation. Comparing the effects of the insect insulin-like proteins and mammalian insulin on growth control and carbohydrate metabolism, those authors suggested that similar cellular mechanisms regulate the development of mammalian pancreatic cells and the insulinproducing cells of D. melanogaster. 5.3
HORMONAL REGULATION OF POLYOL SYNTHESIS DURING COLD-HARDENING
The synthesis of polyols and other cryoprotectants generally appears not to be under hormonal control, but rather is induced directly by low temperature (Storey and Storey, 1991) (Section 4.2.2.1). Although some investigations have demonstrated that developmental hormones, particularly juvenile hormone, affect polyol synthesis, results between studies and different species are often contradictory and a general pattern of hormonal influence is unclear (Zachariassen and Lundheim, 1992). 5.4
DIAPAUSE HORMONE AND TREHALOSE HYDROLYSIS
A special case of hormonal regulation of trehalase occurs during glycogen synthesis in B. mori, where diapause hormone released from the maternal subesophageal ganglion facilitates the synthesis and accumulation of glycogen by oocytes destined for embryonic diapause (Yamashita et al., 1981; Yamashita, 1983; Yamashita and Suzuki, 1991). Glucose is provided through the action of trehalase in the oocytes and follicle cells of the ovary, and haemolymph trehalose hydrolysis, the rate-limiting step in glycogen synthesis, is under direct hormonal control (Ikeda et al., 1993; Su et al., 1994). Injection of trehazolin, a potent trehalase inhibitor, into pupae in the mid-stage of pupal-adult development markedly reduces the accumulation of glycogen in eggs and increases haemolymph trehalose (Katagiri et al., 1998). Following oviposition, the accumulated glycogen in diapausing eggs is quickly broken
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down to glycerol and sorbitol, antifreezes that serve to protect diapausing eggs from damage by cold exposure (see Sections 4.2.2.1 and 7.2.1).
6
Interactions of trehalose and lipid metabolism
Synthesis of trehalose from glycogen by fat body of the cockroach Leucophaea maderae is accompanied by an increase in oxygen consumption associated with fatty acid oxidation (Wiens and Gilbert, 1965). Treatment of American cockroach fat body with extracts of corpora cardiaca decreases glucose oxidation while stimulating fatty acid oxidation, suggesting that the latter provides the energy required for trehalose synthesis (McDougall and Steele, 1988). Indeed, pent-4-enoic acid, an inhibitor of oxidation, reduces trehalose synthesis. Isolated trophocytes from disaggregated fat body of P. americana show increased rates of trehalose synthesis when incubated with a variety of fatty acids, including the polyunsaturates, linoleic and arachidonic acids (Ali and Steele, 1997). Prostaglandin F2 increases trehalose synthesis by trophocytes to a similar degree as hypertrehalosemic hormone. This supports the conclusion that arachidonic acid and eicosenoids synthesized from linoleic acid and released in response to phospholipase A2 action, may play a key role in mediating the actions of hypertrehalosemic hormone (see Section 5.1.1). Several investigations demonstrate that trehalose is important in regulating the release of adipokinetic hormone for mobilizing lipids during flight in migratory locust (Cheeseman et al., 1976; van der Horst et al., 1979; Golding and Pow, 1991). During and immediately following flight initiation, haemolymph trehalose is utilized for energy. While trehalose levels remain high, trehalose, possibly after hydrolysis to glucose, inhibits the release of adipokinetic hormone at the level of the adipokinetic cells in the corpora cardiaca (Passier et al., 1997). As haemolymph trehalose level declines the inhibitory effect is removed and lipids are mobilized to provide energy substrate for flight. Other studies suggest that trehalose also blocks the action of adipokinetic hormone at the level of the fat body, by inhibiting the loading of diglyceride by lipophorin (Lum and Chino, 1990). In adult M. sexta, trehalose decreases fat body triglyceride lipase activity, but does not affect lipoprotein binding (Arrese et al., 1996).
7 7.1
Physiological roles ENERGY STORAGE
That haemolymph trehalose serves as an energy store is clear from investigations of insect flight (see Section 4.2.2). The advantage of trehalose for rapid energy production has a significant cost. From an energetic
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standpoint, trehalose is a less efficient storage carbohydrate than is glycogen. Trehalose synthesis requires more energy than glycogen synthesis for each unit of glucose stored, two units of glucose phosphate rather than one, and upon hydrolysis yields glucose, in contrast to glucose phosphate from glycogen. While haemolymph trehalose may be the major energy store of a few insects, trehalose generally is present at lower amounts than is glycogen or fat. Few investigations, however, have estimated the actual contribution of trehalose to total energy reserves, which undoubtedly varies between species as well as stage of development. 7.2 7.2.1
STRESS PROTECTION
Cryopreservation
Many insects accumulate in their haemolymph and other tissues small molecular weight metabolites and various proteins, that serve to protect the animal from effects of low temperatures that would otherwise prove damaging or lethal (Storey and Storey, 1988; Duman et al., 1991; Lee, 1991). In freezesusceptible insects that avoid freezing and survive temperatures below their normal freezing points, these compounds act as antifreezes to decrease their melting and supercooling points (Somme, 1982; Storey and Storey, 1991). In such cases, supercooling points are typically 10 to 30 C, although they may be much lower, with various metabolites accumulating to concentrations of 0.2 to 5 M (Miller, 1982; Zachariassen, 1985, 1991). The supercooling capacity, the difference between the melting and supercooling points, is a colligative property related to the molal concentration of solute particles affecting water structure (Dick, 1979; Zachariassen, 1991). Glycerol, synthesized principally from glycogen, is the most important and common in this regard, although most polyols share common features of high solubility, non-toxicity and solute compatibility (Storey and Storey, 1991). Many cold-hardy insects accumulate trehalose and/or other sugars, rather than, or in addition to polyols (Storey and Storey, 1991; Kimura et al., 1992; Fields et al., 1998; Ring and Danks, 1998). Trehalose accumulation is particularly well documented for diapausing pupae of the silkmoth, P. cynthia (Hawakawa and Chino, 1981, 1982a,b) (Table 2). The synthesis and metabolism of various low molecular weight antifreezes are described above (see Section 4.2.2.1). Antifreezes may also reduce the supercooling point by inactivating ice nucleators. This effect alone may reduce supercooling points to approximately 20 C (Lee et al., 1981; Duman et al., 1991). In contrast to freeze avoidance, the overwintering strategy of many coldhardy insects involves a tolerance to freezing (Zachariassen, 1985; Storey and Storey, 1988; Lee, 1991). Often the term cryoprotection is applied narrowly to describe this phenomenon only. Here the supercooling point is not decreased,
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but rather is increased in the presence of the same small molecules that serve as antifreeze to reduce the supercooling point in freeze-avoiding insects. This increase in supercooling point ensures that nucleation and ice formation are restricted to the extracellular environment. Only in this manner is freezing and ice formation generally tolerable (Bale, 1996). Regulation of the supercooling point in freeze-tolerant insects involves a variety of ice nucleators including proteins, lipoproteins and ice nucleating bacteria (Duman et al., 1991). As ice forms in the extracellular space and haemolymph, the excluded solutes are concentrated, resulting in an osmotic gradient that removes water from unfrozen tissues and cells thereby causing dehydration (Zachariassen, 1985, 1991). This in turn results in concentration of the intracellular solutes that increases melting point and depresses nucleation temperature, further reducing the potential for intracellular freezing. In freeze-tolerant insects, polyols and sugars act to protect tissues from the effects of osmotic stress and rupture of the cell membrane, as well as fusion of the cell membrane and intracellular membranes (Meryman, 1971, 1974; Taylor, 1987). In addition, protection from potentially harmful effects is afforded due to concentration of intracellular components, particularly ionic species. All polyols and sugars that can be transported across or directly penetrate membranes afford cryoprotection, in part by decreasing the osmotic gradient and cell shrinkage. The presence of cryoprotectants also limits the rate and amount of ice formation in the extracellular space, limiting the concentration of solutes. Perhaps the most important effects of cyroprotectants relate to their effects on membrane stability and protein structure. Considered by many to be the principal mechanisms that afford protection from dehydration, these effects are due to noncolligative properties that vary among cryoprotectants. Extensive investigations have been conducted on other organisms, particularly yeasts and nematodes, many of which undergo similar dehydration upon entering a quiescent state of anhydrobiosis (Crowe et al., 1992; de Araujo, 1996; Behm, 1997; Womersley et al., 1998). Many nematodes, for example, selectively accumulate trehalose that often approaches 20% of their dry weight (Womersley, 1987). Unlike the colligative effects of cryoprotectants, where high levels are required for freeze protection, the effects of cryoprotectants, including trehalose, on membranes and proteins does not necessarily require high concentrations (see for example, Higa and Womersley, 1993). Trehalose is of particular significance and is more effective than other cryoprotectants such as glycerol or sucrose in stabilizing membranes. It reduces membrane fusion, as well as lateral phase separation within membranes due to liquid crystalline to gel phase transitions (Quinn, 1985; Loomis, 1991; Tsonev and Tihova, 1994; Womersley and Higa, 1998). During dehydration, trehalose initially acts as a kosmotropic agent, an ‘order-maker’, stabilizing the structure of water (Sanderson et al., 1991). Gradually trehalose serves to replace water, forming hydrogen bonds directly with proteins and phospholipids, thereby
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maintaining the membrane in the liquid crystalline phase (Crowe et al., 1984, 1987, 1992; Hianik et al., 1996; Hanamura et al., 1998). Lacking intramolecular hydrogen bonds, trehalose dihydrate more readily replaces water than other sugars, conforming to the irregular surfaces of membranes (Panek, 1995; Akao et al., 2001). Last, the resistance of trehalose to acid hydrolysis makes it ideal for replacement of water in proteins (Rosser, 1991; O’Brien, 1996). Conformational analyses demonstrate that trehalose binds to water molecules to produce ice-like clusters similar to those that occur spontaneously between water molecules in pure ice (Akao et al., 1998). As water is removed from a membrane or protein surface, trehalose effectively replaces the water because the spatial arrangement of hydroxyl groups is the same as that of the water cluster – an effect referred to as the ‘water-camouflage effect’. Vitrification is critically important for maintaining the structure of dehydrated membranes (Crowe et al., 1996). In vitrified membranes, the intra- and intermolecular motions necessary for fusion and protein denaturation are hindered or absent (Hagen et al., 1995; Sastry and Agmon, 1997; Cordone et al., 1998). Glass formation is considered by many to be the principal reason for the effectiveness of trehalose in protecting cells and tissues from dehydration (Green and Angell, 1989; Crowe et al., 1998; Mariani et al., 1999; Akao et al., 2001). The dehydration, crystallization and vitrification of trehalose has been examined in some detail (Sussich et al., 1999, 2001; Akao et al., 2001). Overall, trehalose, of all compounds thus far examined, best maintains the native structure of membrane proteins in the dried state under suboptimal conditions (Prestrelski et al., 1993; Uritani et al., 1995; de Araujo, 1996; Crowe et al., 2001).
7.2.2
Protein stabilization during osmotic and thermal stress
Studies with a variety of organisms, including bacteria, yeast, algae, and a few vascular plants, demonstrate that trehalose stabilizes proteins and alleviates osmotic and thermal stress in completely hydrated biological materials, as well as in the dehydrated or desiccated state (Wiemken, 1990; Panek and Panek, 1990; D’Amore et al., 1991; Muller et al., 1995; de Araujo, 1996; Takenaka et al., 1997; Goddijn and van Dun, 1999). In many yeasts, induction of trehalose-6-phosphate synthase and accumulation of trehalose accompanies heat stress (de Virgilio et al., 1990, 1994; Eleutherio et al., 1993; Arguelles, 1997). Similarly, hyperosmotic stress induces accumulation of intracellular trehalose that imparts resistance to otherwise deleterious effects (Dupray et al., 1995; Mikkat et al., 1997; Hounsa et al., 1998; Garcia de Castro and Tunnacliffe, 2000). Trehalose also prevents oxidative stress and the damaging effects of oxygen free radicals (Mouradian et al., 1985; Benaroudj et al., 2001). Prevention of radiation damage to DNA by trehalose has been
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reported, although other disaccharides appear equally protective (Yoshinaga et al., 1997). In contrast to the extensive literature demonstrating the potential role for trehalose in stress protection in yeast and other non-insect organisms, only a few investigations have been conducted with insects. In M. sexta, trehalose prevents thermal denaturation of fat body glycogen phosphorylase b (MeyerFernandez et al., 2000). Glycerol, sorbitol, sucrose, maltose and lactose had a lesser effect or were ineffective. Trehalose at 600 mM in the presence of 20 mM AMP maintains 90% of enzyme activity after 20 min at 60 C. Although trehalose accumulation occurs during high temperature exposure of summer diapausing onion maggot, Delia antiqua, non-diapausing pupae that fail to accumulate trehalose are also high temperature tolerant (Nomura and Ishikawa, 2001). Future studies will undoubtedly demonstrate how trehalose and other metabolites act to protect insects from various stressors. Insects produce heat shock proteins in response to thermal stress (Pauli et al., 1992), and heat shock protein/trehalose interactions are known to affect heat shock protein stability and function in yeast (Singer and Lindquist, 1998a). Induction of trehalose synthesis and accumulation by D. melanogaster increases tolerance to anoxia (Chen et al., 2002). Because anoxia simultaneously induces the synthesis of heat shock proteins, those authors conclude that trehalose interacts with heat shock proteins to minimize protein damage. The bases for stress protection and protein stablization by trehalose under hydrated conditions are incompletely understood. Trehalose has been shown more effective than many other sugars and polyols at protecting various enzymes and structural proteins from heat inactivation and protein aggregation (Hottiger et al., 1994; Sola-Penna and Meyer-Fernandes, 1998). Having a larger hydration volume than other sugars, trehalose occupies a much larger volume and substitutes more water molecules (Sola-Penna and Meyer-Fernandes, 1998). Absence of intramolecular hydrogen bonding ensures that the hydroxyl groups of trehalose interact preferentially with the solvent rather than among themselves (Bonanno et al., 1998). Of the three common disaccharides, trehalose has a greater hydration ability than sucrose or maltose, and more effectively lowers the mobility of water molecules hydrogen bonded with these disaccharides (Kawai et al., 1992; Sakurai et al., 1997). Trehalose maintains proteins in the partially folded state during heat shock, thereby facilitating the refolding by cellular chaperones, including heat shock proteins, when normal conditions prevail (Singer and Lindquist, 1998a,b; Simola et al., 2000). Thermodynamic studies on preferential interaction and thermal unfolding indicate that under conditions where protein is native, trehalose is preferentially bound, while under conditions where protein is denatured and unfolded, trehalose is preferentially excluded (Xie and Timasheff, 1997). Stabilization of the native state is thus achieved by a greater preferential hydration of the denatured than of the native protein structure.
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Despite questions concerning the singularity and universality of the beneficial effects of trehalose (Crowe et al., 1996; Lewis et al., 1997; Womersley et al., 1998), the disaccharide is widely regarded as a major factor in biostabilization and stress protection (van Laere, 1989; de Araujo, 1996; Arguelles, 2000; Tunnacliffe et al., 2001).
7.3
REGULATION OF FEEDING
Many insects offered a variety of foods differing in nutrient content, select among foods, feeding in proportions that satisfy their requirement for a particular balance in the intake of different nutrients. This feeding behaviour, sometimes called dietary self selection, has important ecological significance for understanding how insects interact with plants and other food sources in their environment (Waldbauer and Friedman, 1991). Acting in concert with post-injective physiological effects, changes in digestion and assimilation, self-selection behaviour is often compensatory, allowing insects to avoid otherwise potentially deleterious effects of feeding on nutritionally inadequate foods. Geometric models of nutrient consumption have established that self-selection behaviour achieves a dynamic ‘intake target’, a balance of different nutrients that optimizes growth and development under specific nutritional conditions (Simpson and Raubenheimer, 1993a; Raubenheimer and Simpson, 1999). Choice experiments employing chemically defined diets of varying nutrient composition demonstrate optimal intake ratios for protein and carbohydrate consumption by a variety of insects at different developmental stages under a variety of nutritional conditions. Last instar migratory locusts, for instance, consume a ratio of protein: digestible carbohydrate of approximately 0.7, more carbohydrate than protein, when offered one of several pairs of chemically defined diets, each having the same total nutrient content, but differing in the amounts of protein and carbohydrate (Chambers et al., 1995). On the other hand, last instar larvae of the lepidopteran Helicoverpa zea consume a ratio of 4.0, more protein than carbohydrate, when offered a single pair of chemically defined diets, one with protein but without carbohydrate, and a second with carbohydrate but lacking protein (Waldbauer et al., 1984). The difference reflects the differing needs of the two insects during those stages of development and under those specific nutritional conditions. Regulation of feeding behaviour and nutrient intake involves a complex interaction of physiological processes that integrate information concerning the nutrient composition of food and the nutritional state of the insect (Bernays and Simpson, 1982; Simpson, 1995; Simpson et al., 1995; Simpson and Raubenheimer, 1996). Because haemolymph composition reflects nutrient intake, the presence and concentration of haemolymph metabolites, including trehalose, are indicators of nutritional status that serve as a basis for feedback
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mechanisms regulating food choice and nutrient intake (Simpson and Raubenheimer, 1993b). Studies with several insects establish that feeding is influenced by neural input through the central nervous system from chemoreceptors responding to the presence of nutrients in food (Morita and Shiraishi, 1968; Bernays and Simpson, 1982; Shiraishi and Yano, 1984; Schiff et al., 1989; Simpson and Raubenheimer, 1996). Further, the sensitivity of these receptors is influenced by nutritional status as reflected by haemolymph composition (Evans and Dethier, 1957; Blaney et al., 1986; Simpson et al., 1991, 1995; Amakawa, 2001). Feeding studies with the migratory locust demonstrate that chemosensory responsiveness in taste receptors is regulated by the concentrations of haemolymph amino acids, particularly eight amino acids, detected directly by receptors through contact with the circulating haemolymph (Simpson and Simpson, 1992). Sensitivity increases as haemolymph concentration decreases and vice versa. In the locust, this portion of nutrient feedback does not involve the central nervous system. Modulation of chemoreceptor sensitivity to sugars also occurs, but independently of that for amino acids (Simpson et al., 1990a, 1991). Haemolymph levels of amino acids, trehalose and glucose were positively related to the levels of these nutrients in the diet (Zanotto et al., 1996). Thus, locusts feeding on high protein diets have high haemolymph concentrations of most amino acids, while those feeding on high carbohydrate diets have high levels of both trehalose and glucose. In response to the information feedback provided by haemolymph composition on chemosensory activity, locusts continuously regulate carbohydrate and protein intake (Raubenheimer and Simpson, 1990, 1993). Finally, experiments with locusts conditioned on diets having variable nutrient levels demonstrate that the quantity of diet and/or of nutrients ingested, is regulated by nutritional state in a manner consistent with the effects of conditioning on haemolymph metabolite levels (Simpson, 1995; Simpson et al., 1988, 1990b, 1991). Limited information is available concerning the molecular basis for taste reception and specificity for individual nutrients and other chemicals in insects. However, G protein-coupled receptors and receptor genes specific for trehalose are now known in D. melanogaster (Ishimoto et al., 2000; Dahanukar et al., 2001; Ueno et al., 2001) Studies with lepidopteran larvae also correlate feeding behaviour and nutrient intake with food content in a manner consistent with feedback regulation involving haemolymph composition. Spodoptera littoralis larvae conditioned on a high protein diet, having twice the level of casein as sucrose, subsequently choose a high carbohydrate diet, with twice the amount of sucrose as casein, when offered a choice of both diets (Simpson et al., 1988). Similarly, larvae conditioned on a high sucrose diet, subsequently chose the high protein diet, although they are less sensitive than the locust to prior deprivation of protein than to carbohydrate (Simpson et al., 1990b). As with the locust, the responsiveness of taste chemoreception to dietary amino acids
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and sugars in S. littoralis is influenced by conditioning on diets having variable nutrient composition (Simmonds et al., 1992). Moreover, chemosensory feedback by dietary amino acids and sugar also appears to be mediated independently in S. littoralis. A relationship between haemolymph trehalose concentration and dietary self-selection by H. zea has been established (Friedman et al., 1991). Larvae maintained on a chemically defined diet containing casein but lacking sucrose have very low levels of haemolymph trehalose, while those feeding on a diet with sucrose, but without casein, have trehalose concentrations several-fold higher. When larvae maintained on a high protein or high carbohydrate diet are transferred to the other diet, haemolymph trehalose sharply decreases and increases, respectively. When larvae fed the casein diet are offered a choice of both diets on which to feed, larvae initially feed on the high sucrose diet (Fig. 10). Following a few hours feeding larvae then switch to the casein diet; this change coincides with a rapid increase in haemolymph trehalose concentration due to trehalose synthesis from dietary sucrose. If larvae are injected with trehalose at the time they are offered the dietary choice, they fail to feed selectively on the high sucrose diet. Injection of lactose significantly delays the effect of increased feeding on the sucrose diet, indicating that the haemolymph feedback mechanism may be more specific for trehalose than for other sugars. As with S. littoralis, H. zea larvae appear less sensitive to prior protein deprivation than to carbohydrate deprivation.
FIG. 10 Effect of dietary conditioning on feeding behaviour of 5th instar Helicoverpa zea larvae. Number of larvae feeding on a carbohydrate diet (d) versus a protein diet (s) (Left) after feeding on the high protein diet for 15 h and offered a choice of both diets. (Right) after feeding on the high protein diet for 15 h, injected with 4 l of 0.25 M trehalose and offered a choice of both diets. (Data after Friedman et al., 1991.)
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Studies with M. sexta employing diets with variable levels of casein and sucrose further establish the nature of the feedback between haemolymph trehalose and self-selection behaviour, and demonstrate that dietary nutrient levels interact to affect nutrient intake and haemolymph trehalose (Thompson and Redak, 2000). Larvae fed diets having a variable level of casein, but a constant level of sucrose, display a decrease in carbohydrate consumption as dietary casein level increases, and a decreasing level of haemolymph trehalose (Fig. 11). Larvae fed diets having variable casein, but without sucrose, show increased haemolymph trehalose with increasing dietary casein. When conditioned on these diets, larvae offered a choice of a high protein diet without sucrose and a high carbohydrate diet without casein, feed in a manner related to haemolymph trehalose concentration. Below approximately 30 mM haemolymph trehalose, larvae chose the high carbohydrate diet during the first two hours. Above that trehalose concentration, larvae fed on the high protein diet. Further, larvae with haemolymph trehalose higher than 30 mM were gluconeogenic, and trehalose was maintained by de novo carbohydrate synthesis (Thompson et al., 2002) (see Section 4.1.4.1). Subsequent studies further characterize the relationship between dietary protein and carbohydrate intake, haemolymph sugar level and de novo trehalose synthesis in M. sexta larvae (Thompson et al., 2001, 2003).
FIG. 11 Effect of dietary protein level on haemolymph trehalose concentration and feeding behaviour of 5th instar Manduca sexta larvae. (Left) Haemolymph trehalose level on diets with 12.5 g/l sucrose (d) and on carbohydrate free diets (s). (Right) Percentage of larvae feeding on a high carbohydrate diet after conditioning for 24 h on diets containing 12.5 g/l sucrose (d) and on carbohydrate free diets (s). (Data after Thompson and Redak, 2000.)
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Simpson and Raubenheimer (1993b, 1996) outlined a dynamic model to illustrate how the interaction of food composition, chemosensory responsiveness and central nervous system input affects nutrient intake and haemolymph composition. Based on studies of locusts and lepidopteran larvae, their model considers how nutritional status, reflected by haemolymph composition, acts to mediate those interactions over time. Mechanisms of regulation of self-selection behaviour in response to haemolymph composition are poorly understood in lepidopteran larvae, but appear to involve feedback through the central nervous system. Investigations with isolated nervous system of Mamestra brassicae, for example, demonstrate that specific afferent neurones are chemosensory, responding to elevated levels of trehalose in the haemolymph (Okajima et al., 1989). In this case, stimulation resulted in the release of ecdysone from the prothoracic glands, accelerating development. Additional regulatory mechanisms over self-selection behaviour possibly include hormones, neurohormones and/or biogenic amines. The potential roles of hypertrehalosemic and hypotrehalosemic factors are clear from the above discussion of the effects of these hormones on metabolism and haemolymph sugar composition (see Sections 5.1 and 5.2). Biogenic amines may be involved as well, and nutritional investigations with H. zea implicate serotonin as a regulatory factor. H. zea larvae maintained on a high sucrose diet lacking protein have higher levels of brain serotonin than larvae fed a high casein diet without carbohydrate (Cohen et al., 1988). Larvae treated with p-chlorophenylalanine, an inhibitor of serotonin synthesis, had low brain serotonin levels. When offered a choice of a high protein and a high carbohydrate diet, these larvae consume proportionately more of the carbohydrate diet and less of the protein diet than untreated insects. Although dietary supplements of tryptophan, the amino acid precursor of serotonin, have no effect on brain serotonin level, consumption of the protein diet is proportionately higher in larvae offered the same dietary choice as above. The basis for serotonin action on food selection by H. zea is unexplained, but serotonin concentrations, as well as those of other biogenic amines including dopamine, have long been known to affect and be affected by feeding in mammals (Shor-Posner et al., 1986; Fernstrom and Fernstrom, 1995, 2001; Huether et al., 1998). These biogenic amines also mediate some sensory responses in insects. In M. sexta, for instance, a single serotonin neuron innervates all the glomeruli of the olfactory lobe. High concentrations of serotonin enhance lobe activity while low concentrations reduce activity (Kloppenburg and Hildebrand, 1995). One possible basis for serotonin action concerns alteration of haemolymph metabolite levels through effects on haemolymph volume. Together with a peptide diuretic hormone, serotonin induces diuresis in Rhodnius prolixus following haemolymph feeding (Maddrell et al., 1991). Similar effects occur in larval Aedes aegypti (Clark and Bradley, 1996). If serotonin affects diuresis in insects generally, this may serve as a means for regulating haemolymph
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metabolite concentrations and in turn feeding behaviour. In lepidopteran larvae, including M. sexta, compensatory increases in feeding in response to dietary nutrient dilution are compensated by increased excretion of water (Timmins et al., 1988). How such effects on diuresis influence individual haemolymph nutrients can only be speculated on, but alterations of haemolymph volume might be reflected by overall changes in haemolymph osmolality. Changes in haemolymph osmolality occur in response to variable nutrient intake, and affect feeding responses in migratory locust (Abisgold and Simpson, 1987). Less is known about lepidopteran larvae, although in the case of H. zea, haemolymph osmolality appears unaffected by changes in dietary protein and carbohydrate levels (Friedman et al., 1991). Gut distension and stimulation of gut stretch receptors play a role in regulating food intake in the locust, possibly acting through the release of neurohormones and/or biogenic amines within or from the central nervous system (Simpson et al., 1995). Gut and body wall stretch receptors are also involved in regulation of feeding by M. sexta (Reinecke et al., 1973; Griss et al., 1991; Rowell and Simpson, 1992), but direct feedback from the presence of nutrients in the gut lumen also appears to play a major role (Timmins and Reynolds, 1992). Here the midgut endocrine system is likely involved, possibly affecting assimilation and thereby haemolymph composition. Studies with M. sexta, as well as other insects, have demonstrated a rich peptidergic innervation of the gut (Zitnan et al., 1993, 1995). The contents of these neurons are immunoreactive with antibodies of most mammalian gastroenteropancreatic and other peptides, including FMRFamide-related peptides. These hormones and peptides probably play a role in mediating gut function and possibly regulate digestive enzyme secretion and absorption in response to feedback from the haemolymph, directly or indirectly through the central nervous system. FMRFamines, for example, are known to influence digestion in insects (Elia et al., 1993; Fujisawa et al., 1993; Orchard et al., 2001). Endocrine cells located within the insect midgut epithelium also contain a rich complement of peptides (Sehnal and Zitnan, 1990; Zitnan et al., 1993; Kingan et al., 1997). For example, the insulin-like peptides in the gut regulate fat body glucose metabolism and affect haemolymph trehalose and glucose levels in several insects (see Section 5.2). Although the mechanisms of endocrine cell secretion are unknown, these hormones likely play an important role in the regulation of haemolymph composition and nutrient intake. Studies with the migratory locust demonstrate that the release of FMRFamide-related peptides from midgut endocrine cells is related to feeding and influenced by the nutritional content of food consumed (Zudaire et al., 1998). Further, these endocrine cells are in simultaneous contact with the haemolymph, the urine passing from the malpighian tubules into the gut lumen, as well as the contents of the gut lumen. Noting that these three compartments, the food, haemolymph and excreta, are the key components for integration of nutritional
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homeostasis, Zudaire et al. suggested that these endocrines act as an ‘integrative receptor-secretory’ system.
8
Conclusion
Trehalose, the insect blood sugar, is involved in a multitude of physiological functions. Only during the past few years have some of these roles come to be recognized. Consideration of their potential importance to insects results in large part from studies of non-insect organisms, particularly yeast. When the subject of trehalose in insects was last comprehensively reviewed (Friedman, 1978), trehalose was considered an energy store. This is understandable, because at the time, unravelling the metabolic bases for sustaining flight in various insects was a principal research focus in insect physiology. Despite suggestions that trehalose likely plays a much more complex role in insect physiology than just an energy store (Jungreis, 1980), subsequent treatments continued to consider the function of the disaccharide very narrowly (Friedman, 1985; Downer, 1980; Ziegler, 1984). Recent reviews (Becker et al., 1996) as well as textbooks (Nation, 2002) continue this emphasis: ‘Trehalose . . . serves as a circulating energy source . . .’ (Klowden, 2002). It is interesting and perhaps ironic then that the role of trehalose as an energy store may not be universal. The case has been made in fungi, for instance, that trehalose does not serve as an energy store, although it is present at high levels in cells and spores (van Laere, 1989; Wiemken, 1990). Another historic research focus with insects is the regulation of trehalose metabolism. The hormonal regulation of this metabolism and haemolymph trehalose concentration is no longer in doubt. As the above review attests, insects have many hypertrehalosemic and hypotrehalosemic factors that directly affect trehalose synthesis and degradation. These hormones not only influence haemolymph trehalose level in response to specific physiological stresses, but also act to affect or maintain haemolymph trehalose levels under normal resting conditions. This is clear from ligation experiments or by extirpation of the brain or specific areas of the brain. A consistent pattern of effects, however, is not clear. As described above, for example, removal of the corpora cardiaca of some insects results in a decrease in haemolymph trehalose, presumably due to the absence of some hypertrehalosemic factor, while removal from others produces the opposite effect, suggesting a hypotrehalosemic factor. It may be that different regulatory mechanisms exist in different species and a different hormone or balance of hormones is involved in regulating trehalose metabolism and haemolymph trehalose level. Regarding the effects of cardiacectomy on haemolymph sugar, the results noted above may reflect a difference between lepidopteran and dipteran insects, respectively. Experiments employing hormone injections to confirm the
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effects of ligature or extirpation, or merely to induce effects suggestive of the normal activities of hormones, have not always clarified our understanding. Often, extracts of tissues such as the corpora cardiaca are employed, rather than pure compounds. The corpora cardiaca, as well as other brain tissues, contain many peptides, proteins and other regulatory molecules. These are undoubtedly released in a carefully programmed manner in response to specific physiological cues. Even with studies employing pure compounds then, the significance of results to regulation under normal conditions must be viewed with caution. Metabolic regulation is achieved by a dynamic balance of several hormones and other regulatory molecules. Studies employing molecular approaches, where the expression of genes for several hormones and their receptors can be simultaneously measured under specific physiological conditions, will help provide a fuller understanding of the hormonal regulation of trehalose metabolism in various insect species. Many studies cited above suggest that haemolymph glucose is, in fact, the metabolite acting as a cue to influence hormonal regulation of trehalose synthesis and degradation. Overall, however, the results are very contradictory, with glucose being elevated in one study, while being reduced or unchanged in another, in response to the same or similar physiological condition or treatment. Again, mechanistic differences between insect species may offer an explanation. There is presently little conclusive evidence demonstrating that glucose plays a principal role in the hormonal regulation of trehalose metabolism. What is clear, at this time, is that haemolymph sugar levels are not homeostatically regulated. Indeed, this is essential. Both glucose and trehalose occur at greatly variable levels. To effectively fulfill the roles outlined above, whether energy store or stress protector, trehalose must occur over a wide range of concentrations. Regulation of haemolymph trehalose level is an example of functional conservation through enantiostasis. Several authors cited above, investigating haemolymph trehalose in insects, incorrectly refer to their studies as investigations of homeostasis. This is an implied preference for characterizing physiological stability in the broad manner of C. Bernard. Those authors are not alone in this view. Describing various aspects of invertebrate physiology and metabolism, others have lamented W. B. Canon’s refinement of Claude Bernard’s original concept (Mangum and Towle, 1977; Hochachka and Somero, 1984). Mangum and Towle point out that the refinement obscures subtle but important differences in mechanisms of physiological adaptation. Thus, the distinction between the narrow and the broad views is not trivial. Nevertheless, without the refinement, the utility of the term homeostasis, in reference to a conservation of physiological state within very narrow limits, is lost. Homeostasis is one means, but not the only means, by which physiological stability is achieved. In his thoughtful review titled ‘Trehalose regulation, one aspect of metabolic homeostasis’, Friedman (1978) remarked: ‘To state the problem succinctly the
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concentration of blood trehalose is maintained by a condition of equilibrium between the intake of carbohydrate and synthesis of trehalose on the one hand and metabolic utilization and other loss of this sugar on the other. What forces operate to promote the steady-state condition of the mix?’ Regarding the mechanisms of regulation over trehalose metabolism, we now know the answers to many questions posed by Friedman in that review. Today, the question might be restated as: How do the forces that regulate trehalose metabolism operate and interact with other physiological systems to constantly alter the steady-state concentration of trehalose in response to changing conditions, and how does trehalose concentration in turn affect those same physiological systems? This is the crux to understanding the enantiostatic regulation of blood trehalose.
Acknowledgements The author is grateful to Drs. M. M. Midland and R. M. Wing, Department of Chemistry, University of California, Riverside, for thoughtful discussions on the chemistry of carbohydrates.
References Abisgold, J. D. and Simpson, S. J. (1987). The physiology of compensation by locusts for changes in dietary protein. J. Exp. Biol. 129, 329–346. Abou-Seif, M. A. M., Maier, V., Fuchs, J., Mezger, M., Pfeiffer, E. F. and Bounias, M. (1993). Fluctuations of carbohydrates in haemolymph of honeybee (Apis mellifica) after fasting, feeding and stress. Horm. Metab. Res. 25, 4–8. Akao, K.-I., Okubo, Y., Asakawa, N., Inoue, Y. and Sakurai, M. (2001). Infrared spectroscopic study on the properties of the anyhydrous form II of trehalose. Implications for the functional mechanism of trehalose as a biostabilizer. Carb. Res. 334, 233–241. Akao, K.-I., Okubo, Y., Ikeda, T., Inoue, Y. and Sakurai, M. (1998). Infrared spectroscopic study on the structural property of a trehalose-water complex. Chem. Lett. 759–760. Alaoui, A., Moreau, R. and Gourdoux, L. (1997). Effect of deltamethrin on glucose catabolic pathways in the isolated fat body of adult male Locusta migratoria. Comp. Biochem. Physiol. 116C, 17–21. Aldous, B. J., Auffret, A. D. and Franks, F. (1995). The crystallization of hydrates from amorphous carbohydrates. Cryo-Letters 16, 181–186. Ali, I. and Steele, J. E. (1997). Fatty acids stimulate trehalose synthesis in trophocytes of the cockroach (Periplaneta americana) fat body. Gen. Comp. Endocrinol. 108, 290–297. Ali, I., Finley, C. and Steele, J. E. (1998). Evidence for the participation of arachidonic acid metabolites in trehalose efflux from the hormone activated fat body of the cockroach (Periplaneta americana). J. Insect Physiol. 44, 1119–1126.
262
S. N. THOMPSON
Alumot, E., Lensky, Y. and Holstein, P. (1969). Sugars and trehalase in the reproductive organs and hemolymph of the queen and drone honey bees (Apis mellifica L. Var. ligustica Spin.). Comp. Biochem. Physiol. 28, 1419–1425. Amakawa, T. (2001). Effects of age and blood sugar levels on the proboscis extension of the blow fly Phormia regina. J. Insect Physiol. 47, 195–203. Arguelles, J. C. (1997). Thermotolerance and trehalose accumulation induced by heat shock in yeast cells of Candida albicans. FEMS Microbiol. Lett. 146, 65–71. Arguelles, J. C. (2000). Physiological roles of trehalose in bacteria and yeast: a comparative analysis. Arch. Microbiol. 174, 217–224. Arrese, E. L., Rojas-Rivas, B. I. and Wells, M. A. (1996). The use of decapitated insects to study lipid mobilization in adult Manduca sexta: effects of adipokinetic hormone and trehalose on fat body lipase activity. Insect Biochem. Molec. Biol. 26, 775–782. Arslan, A., Standifer, L. N. and Don, H. (1986). Carbohydrates in honey bee hemolymph. Comp. Biochem. Physiol. 84B, 363–367. Azuma, M. and Yamashita, O. (1985). Cellular localization and proposed function of midgut trehalase in the silkworm larva, Bombyx mori. Tiss. Cell 17, 539–551. Bailey, E. (1975). Insect Biochemistry and Function. Part 2 Fuel Supply. In: Biochemistry of insect flight (eds. Candy, D. J., and Kilby, B. A.), pp. 89–176. New York: Chapman and Hall. Bailey, E. and Walker, P. R. (1969). A comparison of the properties of the pyruvate kinases of the fat body and flight muscle of the adult male desert locust. Biochem. J. 111, 359–364. Bale, J. S. (1996). Insect cold hardiness: a matter of life and death. Eur. J. Entomol. 93, 369–382. Bayer, R. (1968). Untersuchungen am kreislaufsystem der wanderheuschrecke (Locusta migratorioides R. et F., Orthorpteroidea). mit besonderer Berucksichtigung des blutdruckes. Zeit. Vergl. Physiol. 58, 76–155. Becker, A. and Wegener, G. (1998). Hypertrehalosaemic neuropeptides decrease levels of the glycolytic signal fructose 2,6-bisphosphate in cockroach fat body. J. Exp. Biol. 201, 1939–1946. Becker, A., Liewald, J. F. and Wegener, G. (1998). Signal transduction in isolated fat body from the cockroach Blaptica dubia exposed to hypertrehalosaemic neuropeptide. J. Comp. Physiol. 168B, 159–167. Becker, A., Liewald, J. F., Stypa, H. and Wegener, G. (2001). Antagonistic effects of hypertrehalosemic neuropeptide on the activities of 6-phosphofructo-1-kinase and fructose-1,6-bisphosphatase in cockroach fat body. Insect Biochem. Molec. Biol. 31, 381–392. Becker, A., Schloder, P., Steele, J. E. and Wegener, G. (1996). The regulation of trehalose metabolism in insects. Experientia 52, 433–439. Bedford, J. J. (1977). The carbohydrate levels of insect hemolymph. Comp. Biochem. Physiol. 57A, 83–86. Behm, C. A. (1997). The role of trehalose in the physiology of nematodes. Int. J. Parasitol. 27, 215–229. Benaroudj, N., Lee, D. H. and Goldberg, A. L. (2001). Trehalose accumulation during cellular stress protects cells and cellular proteins from damage by oxygen radicals. J. Biol. Chem. 276, 24261–24267. Ben Khay, A., Gourdoux, L., Moreau, R. and Dutrieu, J. (1987). Effects of intestinal insulin-like peptide on glucose catabolism in male adult Locusta migratoria. Arch. Insect Biochem. Physiol. 4, 233–239. Berg, J. M., Tymoczko, J. L. and Stryer, L. (2002). Biochemistry. New York: W.H. Freeman.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
263
Bernard, C. (1865). An Introduction to the Study of Experimental Medicine. Green Dover. New York: Trans. H.C. Bernays, E. A. and Chapman, R. F. (2001). Electrophysiological responses of taste cells to nutrient mixtures in the polyphagous caterpillar of Grammia geneura. J. Comp. Physiol. 187A, 205–213. Bernays, E. A. and Simpson, S. J. (1982). Control of food intake. Adv. Insect Physiol. 16, 59–118. Berthelot, M. (1858). Sur le trehalose, nouvelle espece de sucre. Compt. Rend. Hebd. Seanc. Acad. Sci. 46, 1276–1279. Birch, G. G. (1963). Trehaloses. Adv. Carb. Chem. 18, 201–225. Blaney, W. M., Schoonhoven, W. M. and Simmonds, M. S. J. (1986). Sensitivity variations in insect chemoreceptors: a review. Experientia 42, 13–19. Blatt, J. and Roces, F. (2001). Haemolymph sugar levels in foraging honeybees (Apis mellifera carnica): dependence on metabolic rate and in vivo measurement of maximal rates of trehalose synthesis. J. Exp. Biol. 204, 2709–2716. Blatt, J. and Roces, F. (2002a). The control of the proventriculus in the honeybee (Apis mellifera carnica L.) I. A dynamic process influenced by food quality and quantity? J. Insect Physiol. 48, 643–654. Blatt, J. and Roces, F. (2002b). The control of the proventriculus in the honeybee (Apis mellifera carnica L.) II. Feedback mechanisms. J. Insect Physiol. 48, 683–691. Blau, C. and Wegener, G. (1994). Metabolic integration in locust flight: the effect of octopamine on the glycolytic activator fructose 2,6 bisphosphate in flight muscle in vivo. J. Comp. Physiol. 164B, 11–15. Blau, C., Wegener, G. and Candy, D. J. (1994). The effect of octopamine on the glycolytic activator fructose 2,6 bisphosphate in perfused locust flight muscle. Insect Biochem. Molec. Biol. 245, 677–683. Bock, K. and Pedersen, C. (1983). Carbon-13 nuclear magnetic resonance spectroscopy of monosaccharides. Adv. Carb. Chem. Biochem. 41, 27–66. Boiteux, A., Hess, B. and Sel’kov, E. E. (1980). Creative functions of instability and oscillations in metabolic systems. Curr. Top. Cell. Reg. 17, 171–203. Bolander, F. F. (1994). Molecular Endocrinology. New York: Academic Press. Bonanno, G., Noto, R. and Fornili, S. L. (1998). Water interaction with , trehalose: molecular dynamics simulation. J. Chem. Soc. Faraday Trans. 94, 2755–2762. Bounias, M. (1981). A comparison of haemolymph levels and inter-relations of trehalose glucose and fructose in workerbees from different races and hybrids. Comp. Biochem. Physiol. 69B, 471–477. Bounias, M. and Morgan, M. R. J. (1984). Induction of honeybee haemolymph sucrase activity by high levels of dietary sucrose. Comp. Biochem. Physiol. 79B, 75–80. Bounias, M., Moreau, R. and Gourdoux, L. (1986). Effects of honeybee insulinimmunoreactive peptide on haemolymph lipid and carbohydrate. Insect Biochem. 16, 721–731. Bounias, M., Bahjou, A., Gourdoux, L. and Moreau, R. (1993). Molecular activation of a trehalase purified from the fat body of a coleopteran insect (Tenebrio molitor), by an endogenous insulin-like peptide. Biochem. Molec. Biol. Int. 31, 249–266. Bowers, W. S. and Friedman, S. (1963). Mobilisation of fat body glycogen by an extract of corpus cardiacum. Nature, London 198, 685. Brogiolo, W., Stocker, H., Ikeya, T., Rintelen, F., Fernandez, R. and Hafen, E. (2001). An evolutionarily conserved function of the Drosophila insulin receptor and insulinlike peptides in growth control. Curr. Biol. 11, 213–221. Bursell, E. (1981). The role of proline in energy metabolism. In: Energy Metabolism in Insects (ed. Downer, R. G. H.), Chap. 5, pp. 135–154. New York: Plenum.
264
S. N. THOMPSON
Candy, D. J. and Kilby, B. A. (1961). The biosynthesis of trehalose in the locust fat body. Biochem. J. 78, 531–536. Candy, D. J. (1989). Utilization of fuels by the flight muscles. In: Insect Flight (eds. Goldsworthy, G. J., and Wheeler, C. H.), pp. 305–319. Boca Raton: CRC Press. Candy, D. J., Becker, A. and Wegener, G. (1997). Coordination and integration of metabolism in insect flight. Comp. Biochem. Physiol. 117B, 497–512. Candy, D. J. (1978). The regulation of locust flight muscle metabolism by octopamine and other compounds. Insect Biochem. 8, 177–181. Cannon, W. B. (1932). The Wisdom of the Body. New York: Norton. Cao, C. and Brown, M. R. (2001). Localization of an insulin-like peptide in brains of two flies. Cell Tiss. Res. 304, 317–321. Chambers, P. G., Simpson, S. J. and Raubenheimer, D. (1995). Behavioural mechanisms of nutrient balancing in Locusta migratoria nymphs. Anim. Behav. 50, 1513–1523. Chapman, R. F. (1998). The Insects, Structure and Function. pp. 346. New York: Cambridge University Press. Cheeseman, P., Jutsum, A. R. and Goldsworthy, G. J. (1976). Quantitative studies on the release of locust adipokinetic hormone. Physiol. Entomol. 1, 115–121. Chen, A. C. and Friedman, S. (1977a). Hormonal regulation of trehalose metabolism in the blowfly, Phormia regina: interaction between hypertrehalosemic and hypotrehalosemic hormones. J. Insect Physiol. 23, 1223–1232. Chen, A. C. and Friedman, S. (1977b). Hormonal regulation of trehalose metabolism in the blowfly Phormia regina Meig.: effects of cardiacectomy and allatectomy at the subcellular level. Comp. Biochem. Physiol. 58B, 339–344. Chen, Q., Ma, E., Behar, K. L., Xu, T. and Haddad, G. G. (2002). Role of trehalose phosphate synthase in anoxia tolerance and development in Drosophila melanogaster. J. Biol. Chem. 277, 3274–3279. Chen, P. S. (1985). Amino acid and protein metabolism. In: Comparative Insect Physiology, Biochemistry and Pharmacology (eds. Kerkut, G. A., and Gilbert, L. I.), pp. 177–217. New York: Pergamon, Vol. 10. Cherbas, L. and Cherbas, P. (1970). Distribution and metabolism of -ecdysone in pupae of the silkworm Antheraea polyphemus. Biol. Bull. 138, 115–128. Chino, H. (1961). Enzymatic pathways in the formation of sorbitol and glycerol in the diapausing egg of the silkworm, Bombyx mori. II. On the phosphatases. J. Insect Physiol. 6, 231–240. Chino, H. (1963). Respiratory enzyme system of the Bombyx silkworm egg in relation to the mechanism of the formation of sugar alcohols. Arch. Biochem. Biophys. 102, 400–415. Clark, T. M. and Bradley, T. J. (1996). Stimulation of Malpighian tubules from larval Aedes aegypti by secretagogues. J. Insect Physiol. 42, 593–602. Clegg, J. S. and Evans, D. R. (1961). The physiology of blood trehalase and its function during flight in the blowfly. J. Exp. Biol. 38, 771–792. Clifford, K. H. (1980). Stereochemistry of the hydrolysis of trehalose by the enzyme trehalase prepared from the flesh fly Sarcophaga barbata. Eur. J. Biochem. 106, 337–340. Cohen, R. W., Friedman, S. and Waldbauer, G. P. (1988). Physiological control of nutrient self-selection in Heliothis zea larvae: the role of serotonin. J. Insect Physiol. 34, 935–940. Coon, B. F. (1944). Effect of paralytic insecticides on heart pulsations and circulation of blood in the American cockroach. J. Econ. Entomol. 37, 785–789. Cordes, E. H. and Jencks, W. P. (1962). On the mechanism of Schiff base formation and hydrolysis. J. Amer. Chem. Soc. 84, 832–837.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
265
Cordone, L., Galajda, P., Vitrano, E., Gassmann, A., Ostermann, A. and Parak, F. (1998). A reduction of protein specific motions in co-ligated myoglobin embedded in a treahalose glass. Eur. Biophys. J. 27, 173–176. Coxon, B. (1980). Carbon-13 nuclear magnetic resonance spectroscopy of food-related disaccharides and trisaccharides. In: Developments in Food Carbohydrate-2 (ed. Lee, C. K.), pp. 351–390. London: Applied Science. Crabtree, B. and Newsholme, E. A. (1975). Comparative aspects of fuel utilization and metabolism by muscle. In: Insect Muscle (ed. Usherwood, P. N. R.), pp. 405–500. New York: Academic Press. Crowe, J. H., Carpenter, J. F. and Crowe, L. M. (1998). The role of vitrification in anhydrobiosis. Ann. Rev. Physiol. 6, 73–103. Crowe, J. H., Crowe, L. M., Carpenter, J. F. and Wistrom, C. A. (1987). Stabilization of dry phospholipid bilayers and proteins by sugars. Biochem. J. 242, 1–10. Crowe, J. H., Crowe, L. M., Oliver, A. E., Tsvetkova, N., Wolkers, W. and Tablin, F. (2001). The trehalose myth revisited: introduction to a symposium on stabilization of cells in the dry state. Cryobiology 43, 89–105. Crowe, J. H., Hoekstra, F. A. and Crowe, L. M. (1992). Anhydrobiosis. Ann. Rev. Physiol. 54, 579–599. Crowe, L. M., Mouradian, R., Crowe, J. H., Jackson, A. and Womersley, C. (1984). Effects of carbohydrates on membrane stability at low water activities. Biochim. Biophys. Acta 769, 141–150. Crowe, L. M., Reid, D. S. and Crowe, J. H. (1996). Is trehalose special for preserving dry biomaterials? Biophys. J. 71, 2087–2093. D’Amore, T., Crumplen, R. and Stewart, G. G. (1991). The involvement of trehalose in yeast stress tolerance. J. Indust. Microbiol. 7, 191–196. Dadd, R. H. (1985). Nutrition: organisms. In: Comparative Insect Physiology, Biochemistry and Pharmacology, Vol. 4 (eds. Kerkut, G. A., and Gilbert, L. I.), pp. 313–390. New York: Pergamon Press. Dahanukar, A., Foster, K., van der Goes van Naters, W. M. and Carlson, J. R. (2001). A Gr receptor is required for response to the sugar trehalose in taste neurons of Drosophila. Nature Neurosc. 4, 1182–1186. Dahlman, D. L. (1973). Starvation of the tobacco hornworm, Manduca sexta. 1. Changes in hemolymph characteristics of 5th-stage larvae. Ann. Entomol. Soc. Amer. 66, 1023–1029. Dale, J. E. and Sutcliffe, J. F. (1986). Water and solutes in plants. In: Plant physiology, Vol. IX (eds. Steward, J. F., Sutcliffe, J. E., and Dale, J. E.), pp. 455–549. New York: Academic Press. Davenport, A. P. and Evans, P. D. (1984). Stress-induced changes in the octopamine levels of insect haemolymph. Insect Biochem. 14, 135–143. de Araujo, P. S. (1996). The role of trehalose in cell stress. Braz. J. Med. Biol. Res. 29, 873–875. Denlinger, D. L. (1991). Relationship between cold hardiness and diapause. In: Insects at Low Temperature (eds. Lee, R. E., and Dennlinger, D. L.), pp. 174–198. London: Chapman and Hall. de Virgilio, C, Simmen, U., Hottiger, T., Boller, T. and Wiemken, A. (1990). Heat shock induces enzymes of trehalose metabolism, trehalose accumulation, and thermotolerance in Schizosaccharomyces pombe, even in the presence of cycloheximide. FEBS Lett. 273, 107–110. de Virgilio, C., Hottiger, T., Dominguez, J., Boller, T. and Wiemken, A. (1994). The role of trehalose synthesis for the acquisition of thermotolerance in yeast 1. genetic evidence that trehalose is a thermoprotectant. Eur. J. Biochem. 219, 179–186.
266
S. N. THOMPSON
Dick, D. A. (1979). Structure and properties of water in the cell. In: Mechanisms of Osmoregulation in Animals (ed. Giles, R.), pp. 3–45. New York: John Wiley. Dow, J. A. T. (1986). Insect midgut function. Adv. Insect Physiol. 19, 187–327. Dowd, M. K. and Reilly, P. J. (1992). Conformational analysis of trehalose disaccharides and analogues using MM3. J. Comput. Chem. 13, 102–114. Downer, R. G. H. (1979). Induction of hypertrehalosemia by excitation in Periplaneta americana. J. Insect Physiol. 25, 59–63. Downer, R. G. H. and Matthews, J. R. (1977). Production and utilization of glucose in the American cockroach, Periplaneta americana. J. Insect Physiol. 23, 1429–1435. Downer, R. H. G. (1980). Short term hypertrehalosemia induced by octopamine in the American cockroach, Periplaneta americana. In: Insect Neurobiology and Pesticide Actions (Neurotox 79). pp. 335–339. London: Society of Chemical Industry. Duman, J. G., Xu, L., Neven, L. G., Tursman, D. and Wu, D. W. (1991). Hemolymph proteins involved in insect subzero-temperature tolerance: ice nucleators and antifreeze proteins. In: Insects at low temperature (eds. Lee, R. E., and Denlinger, D. L.), pp. 94–127. New York: Chapman and Hall. Dupray, E., Derrien, A. and Pichon, R. (1995). Osmoregulation by trehalose synthesis in Salmonella manhattan after exposure to waste water. Lett. Appl. Microbiol. 20, 148–151. Duve, H. (1978). The presence of a hypoglucemic and hypotrehalocemic hormone in the neurosecretory system of the blowfly Calliphora erythrocephala. Gen. Comp. Endocrinol. 36, 102–110. Egorova, T. A. and Khomidov, Kh. S. (1991). Purification and properties of membranebound trehalase from silkworm eggs. Biochemistry (Moscow) 56, 958–965. Elbein, A. D. (1974). The metabolism of , -trehalose. Adv. Carb. Chem. Biochem. 30, 227–256. Eleutherio, E. C., Araujo, P. S. and Panek, A. D. (1993). Protective role of trehalose during heat stress in Saccharomyces cerevisiae. Cryobiology 30, 591–596. Elia, A. J., Tebrugge, V. A. and Orchard, I. (1993). The pulsatile appearance of FMRFamide-related peptides in the haemolymph and loss of FMRFamide-like immunoreactivity from the neurohaemal areas of Rhodnius prolixus following a blood meal. J. Insect Physiol. 39, 459–469. Eliel, E. L. (1962). Stereochemistry of Carbon Compounds. pp. 266. New York: McGraw-Hill. Evans, D. R. and Dethier, V. G. (1957). The regulation of taste thresholds for sugars in the blow fly. J. Insect Physiol. 1, 3–17. Fan, T. W.-M. (1996). Metabolite profiling by one- and two-dimensional NMR analysis of complex mixtures. Prog. Nuc. Mag. Res. Spec. 28, 161–219. Fell, R. D. (1990). The qualitative and quantitative analysis of insect hemolymph sugars by high performance thin-layer chromatography. Comp. Biochem. Physiol. 95A, 539–544. Fernstrom, J. D. and Fernstrom, M. H. (2001). Monoamines and protein intake: are control mechanisms designed to monitor a threshold intake or a set point? Nutrition Reviews 59, S60–S68. Fernstrom, M. H. and Fernstrom, J. D. (1995). Brain tryptophan concentrations and serotonin synthesis remain responsive to food consumption after the ingestion of sequential meals. Amer. J. Clin. Nutr. 61, 312–319. Ferreira, J. C., Paschoalin, V. M. R., Panek, A. D. and Trugo, L. C. (1997). Comparison of three different methods for trehalose determination in yeast extracts. Food Chem. 60, 251–254.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
267
Fields, P. G., Fleurat-Lessard, F., Lavenseau, L., Febvay, G., Peypelut, L. and Bonnot, G. (1998). The effect of cold acclimation and deacclimation on cold tolerance, trehalose and free amino acid levels in Sitophilus granarius and Cryptolestes ferrugineus (Coleoptera). J. Insect Physiol. 44, 955–965. Fillinger, S., Chaveroche, M.-K., van Dijck, P., de Vries, R., Ruijter, G., Thevelein, J. and d’Enfert, C. (2001). Trehalose is required for the acquisition of tolerance to a variety of stresses in the filamentous fungus Aspergillus nidulans. Microbiology 147, 1851–1862. Florkin, M. and Jeuniaux, C. (1974). Hemolymph composition. In: The Physiology of Insecta, Vol. 6 (ed. Rockstein, M.), pp. 256–307. New York: Academic Press. Fried, B. and Sherma, J. (1999). Thin-layer Chromatography. New York: Marcel Dekker. Friedman, S. (1971). Interactions among sites responsible for trehalose 6-phosphate and trehalose-activated glucose 6-phosphate hydrolysis on trehalose phosphatase isolated from Phormia regina. J. Biol. Chem. 246, 4122–4130. Friedman, S. (1978). Trehalose regulation, one aspect of metabolic homeostasis. Ann. Rev. Entomol. 23, 389–407. Friedman, S. (1985). Carbohydrate metabolism. In: Comparative Insect Physiology, Biochemistry and Pharmacology, Vol. 10 (eds. Kerkut, G. A., and Gilbert, L. I.), pp. 43–76. New York: Pergamon. Friedman, S. and Hsueh, T.-F. (1979). Insect trehalose-6-phosphatase: the unactivated type, as illustrated in Periplaneta americana, and a survey of the ordinal distribution of the two presently known types. Comp. Biochem. Physiol. 64B, 339–344. Friedman, S., Waldbauer, G. P., Eertmoed, J. E., Naeem, M. and Ghent, A. W. (1991). Blood trehalose levels have a role in the control of dietary self-selection by Heliothis zea larvae. J. Insect Physiol. 37, 919–928. Fujisawa, Y., Shimoda, M., Kiguchi, K., Ichikawa, T. and Fujita, N. (1993). The inhibitory effect of a neuropeptide ManducaFLRFamide, on the midgut activity of the sphingid moth, Agricus convolvulis. Zool. Sci. 10, 773–777. Ga¨de, G. (1990). The adipokinetic hormone/red pigment-concentrating hormone peptide family: structures, interrelationships and functions. J. Insect Physiol. 36, 1–12. Ga¨de, G. (1991). Hyperglycaemia or hypertrehalosaemia? The effect of insect neuropeptides on haemolymph sugars. J. Insect Physiol. 37, 483–487. Ga¨de, G., Reynolds, S. E. and Beeching, J. R. (1994). Perspectives in Comparative Endocrinology (eds. Davey, K. G., Peter, R. E., and Tobe, S. S.), pp. 119–128. Ottawa: National Research Council of Canada. Garcia de Castro, A. and Tunnacliffe, A. (2000). Intracellular trehalose improves osmotolerance but not desiccation tolerance in mammalian cells. FEBS Lett. 487, 199–202. Gies, A., Fromm, T. and Ziegler, R. (1988). Energy metabolism in starving larvae of Manduca sexta. Comp. Biochem. Physiol. 91A, 549–555. Gilby, A. R., Wyatt, S. S. and Wyatt, G. R. (1967). Trehalases from the cockroach, Blaberus discoidalis: activation, solubilization and properties of the muscle enzymes and some properties of the intestinal enzyme. Acta Biochim. Polon. 14, 83–100. Goddijn, O. J. M. and van Dun, K. (1999). Trehalose metabolism in plants. TIPS 4, 315–319. Golding, D. W. and Pow, D. V. (1991). Neurosecretion by synaptic terminals in the locust corpus cardiacum: is non-synaptic exocytosis part of the regulated or the constitutive pathway? Biol. Cell 73, 157–162.
268
S. N. THOMPSON
Goldsworthy, G. J. and Ga¨de, G. (1983). The chemistry of hypertrehalosemic factors. In: Endocrinology of Insects (eds. Downer, R. G. H., and Laufer, H.), pp. 109–119. New York: Alan R. Liss. Gourdoux, L. (1980). The catabolism of glucose in Tenebrio molitor: the effects of corpora cardiaca. J. Insect Physiol. 26, 729–738. Gourdoux, L., Lequellec, Y., Moreau, R. and Dutrieu, J. (1983). Gluconeogenesis from some amino acids and its endocrine modification in Tenebrio molitor. Comp. Biochem. Physiol. 74B, 273–276. Gourdoux, L., Lequellec, Y., Puiroux, J. and Moreau, R. (1989). Differential hormonal control of hemolymph carbohydrates in Pieris brassicae L. Comp. Biochem. Physiol. 93, 543–548. Gourdoux, L., Moreau, R., Ben Khay, A. and Dutrieu, J. (1985). In vivo glucose catabolism in male adult Locusta migratoria: effects of corpora cardiaca. Comp. Biochem. Physiol. 81B, 485–489. Green, J. L. and Angell, C. A. (1989). Phase relations and vitrification in saccharidewater solutions and the trehalose anomaly. J. Phys. Chem. 93, 2880–2882. Griss, C., Simpson, S. J., Rohrbacher, J. and Rowell, C. H. F. (1991). Localization in the central nervous system of larval Manduca sexta (Lepidoptera: Sphingidae) of areas responsbile for aspects of feeding behaviour. J. Insect Physiol. 37, 477–482. Grob, R. L. (1995). Modern Practice of Gas Chromatography. New York: John Wiley. Groen, A. K., Vervoorn, R. C., van der Meer, R. and Tager, J. M. (1983). Control of gluconeogenesis in rat liver cells. I. kinetics of the individual enzymes and the effect of glucagon. J. Biol. Chem. 258, 14346–14353. Gussin, A. E. S. and Wyatt, G. R. (1965). Membrane-bound trehalase from cecropia silkmoth muscle. Arch. Biochem. Biophys. 112, 626–634. Hagen, S. J., Hofrichter, J. and Eaton, W. A. (1995). Protein reaction kinetics in a room temperature glass. Science 269, 959–962. Hallsworth, J. E. and Magan, N. (1997). A rapid HPLC protocol for detection of polyols and trehalose. J. Microbiol. Meth. 29, 7–13. Hanamura, T., Asakawa, N., Inoue, Y. and Sakurai, M. (1998). A solid-state 31P NMR study of the interaction between trehalose and DPPC bilayer. Chem. Lett. 713–714. Hanbury, D. (1859). Note on two insect-products from Persia. J. Proc. Linn. Soc. Zool. 3, 178–183. Hansen, K. (1966). Zur cytologischen lokalisation der trehalase in der indireckten flugmuskulatur der insekten. Biochem. Z. 344, 15–25. Harrison, J. F. (2001). Insect acid-base physiology. Annu. Rev. Entomol. 46, 221–250. Hawakawa, Y. and Chino, H. (1981). Temperature-dependent interconversion between glycogen and trehalose in diapausing pupae of Philosamia cynthis Ricini and Pryeri. Insect Biochem. 11, 43–47. Hawakawa, Y. and Chino, H. (1982a). Temperature-dependent activation or inactivation of glycogen phosphorylase and synthase of fat body of the silkworm Philosamia cynthia: the possible mechanism of the temperature-dependent interconversion between glycogen and trehalose. Insect Biochem. 12, 361–366. Hawakawa, Y. and Chino, H. (1982b). Phosphofructokinase as a possible key enzyme regulating glycerol or trehalose accumulation in diapausing insects. Insect Biochem. 12, 639–642. Hawakawa, Y., Jahagirdar, A. P., Yaguchi, M. and Downer, R. G. H. (1989). Purification and characterization of trehalase inhibitor from the hemolymph of the American cockraoch, Periplaneta americana. J. Biol. Chem. 264, 16165–16169. Hayes, D. K., Bird, T. G., Mills, G. D. and Frankoff, H. (1990). Circadian rhythm of trehalose in the face fly Musca autumnalis de Geer. Chronobiol. Int. 7, 413–418.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
269
Heinrich, B. (1993). The Hot-Blooded Insects. pp. 238–241. Cambridge: Harvard University Press. Hianik, T., Dlugopolsky, J., Gyeppessova, M., Sivak, B., Tien, H. T. and OttovaLeitmannova, A. (1996). Stabilization of bilayer lipid membranes on solid supports by trehalose. Bioelectrochem. Bioenerg. 39, 299–302. Higa, L. M. and Womersley, C. Z. (1993). New insights into the anhydrobiotic phenomenon: the effects of trehalose content and differential rates of evaporative water loss on the survival of Aphelenchus avenae. J. Exp. Zool 267, 120–129. Hochachka, P. W. and Somero, G. N. (1984). Biochemical Adaptation. Princeton: Princeton University Press. Holden, C. P. and Storey, K. B. (1993). Regulation of phosphofructokinase and the control of cryoprotectant synthesis in a freeze-avoiding insect. Can. J. Zool. 71, 1895–1899. Holden, C. P. and Storey, K. B. (1995). Fructose-1,6-bisphosphatase from a cold-hardy insect: control of cryoprotectant glycerol catabolism. Arch. Insect Biochem. Physiol. 28, 225–235. Honda, S. (1984). High performance liquid chromatography of mono- and oligosaccharides. Anal. Biochem. 140, 1–47. Horie, Y., Nakasone, S. and Ito, T. (1968). The conversion of 14C-carbohydrates into CO2 and lipid by the silkworm, Bombyx mori. J. Insect Physiol. 14, 971–981. Hottiger, T., de Virgilio, C., Hall, M. N., Boller, T. and Wiemken, A. (1994). The role of trehalose synthesis for the acquisition of thermotolerance in yeast. II. Physiological concentrations of trehalose increase the thermal stability of proteins in vitro. Eur. J. Biochem. 219, 187–193. Hounsa, C.-G., Brandt, E. V., Thevelein, J., Hohmann, S. and Prior, B. A. (1998). Role of trehalose in survival of Saccharomyces cerevisiae under osmotic stress. Microbiology 114, 671–680. Howden, G. F. and Kilby, B. A. (1956). Trehalose and trehalase in the locust. Chem. Ind. 1453–1454. Huether, G., Schmidt, S. and Ruther, E. (1998). Nutritional effects on central serotonergic activity: a hypothesis on the unconscious self-manipulation of mood by food intake and dietary selection. Nutr. Neurosc. 1, 3–7. Iida, K. and Kajiwara, M. (1990). Isolation of the insect metabolite trehalose by highperformance liquid chromatography. J. Chromatog. 515, 573–575. Ikeda, M., Su, Z., Saito, H., Imai, K., Yukihiro, S., Isobe, M. and Yamashita, O. (1993). Induction of embryonic diapause and stimulation of ovary trehalase activity in the silkworm, Bombyx mori, by synthetic diapause hormone. J. Insect Physiol. 39, 889–895. Ishimoto, H., Matsumoto, A. and Tanimura, T. (2000). Molecular identification of a taste receptor gene for trehalose in Drosophila. Science 289, 116–119. Iwami, M. (1990). The genes encoding bombyxin, a brain secretory peptide of Bombyx mori: structure and expression. In: Molting and Metamorphosis (eds. Ohnishi, E., and Ishizaki, H.), pp. 49–66. Berlin: Springer-Verlag. Iwami, M. (2000). Bombyxin: an insect brain peptide that belongs to the insulin family. Zool. Sci. 17, 1035–1044. Iwami, M., Tanaka, A., Hano, N. and Sakurai, S. (1996). Bombyxin gene expression in tissues other than brain detected by reverse transcription-polymerase chain reaction (RT-PCR) and in situ hydridization. Experientia 52, 882–886. Jahagirdar, A. P., Downer, R. G. H. and Viswanatha, T. (1984). Influence of octopamine on trehalase activity in muscle and hemolymph of the American cockroach, Periplaneta americana L. Biochim. Biophys. Acta. 801, 177–183.
270
S. N. THOMPSON
Jahagirdar, A. P., Downer, R. G. H. and Viswanatha, T. (1990). Purification and characterization of trehalose-hydrolyzing enzymes from the thoracic musculature of the American cockroach, Periplaneta americana. Insect Biochem. 20, 511–516. Jencks, W. P. (1959). Studies on the mechanism of oxime and semicarbazone formation. J. Amer. Chem. Soc. 81, 475–481. Joanisse, D. R. and Storey, K. B. (1994a). Enzyme activity profiles in an overwintering population of freeze-tolerant larvae of the gall fly, Eurosta solidaginis. J. Comp. Physiol. 164B, 247–255. Joanisse, D. R. and Storey, K. B. (1994b). Mitochondrial enzymes during overwintering in two species of cold-hardy gall insects. Insect Biochem. Molec. Biol. 24, 145–150. Jones, J. C. (1977). The Circulatory System of Insects. pp. 78–83. Springfield: Charles C. Thomas. Jungreis, A. M. (1980). Hemolymph as a dynamic tissue. In: Insect Biology in the Future (eds. Locke, M., and Smith, D. S.), pp. 273–294. New York: Academic Press. Kageyama, T. (1976). Pathways of carbohydrate metabolism in the eggs of the silkworm, Bombyx mori. Insect Biochem. 6, 507–511. Katagiri, N., Ando, O. and Yamashita, O. (1998). Reduction of glycogen in eggs of the silkworm, Bombyx mori, by use of a trehalose inhibitor, trehazolin, and diapause induction in glycogen-reduced eggs. J. Insect Physiol. 44, 1205–1212. Katz, J. and Wood, H. G. (1960). The use of glucose-C14 for the evaluation of pathways of glucose metabolism. J. Biol. Chem. 235, 2165–2177. Katz, J. and Wood, H. G. (1963). The use of C14O2 yields from glucose-1- and -6-C14 for the evaluation of the pathways of glucose metabolism. J. Biol. Chem. 238, 517–523. Kaufman, M. G., Walker, E. D., Odelson, D. A. and Klug, M. J. (2000). Microbial community ecology and insect nutrition. Amer. Entomol. 46, 173–184. Kawai, H., Sakurai, M., Inoue, Y., Chujo, R. and Kobay Ashi, S. (1992). Hydration of oligosaccharides: anomalous hydration ability of trehalose. Cryobiology 29, 599–606. Keeley, L. L., Park, J. H., Lu, K.-H. and Bradfield, J. Y. (1996). Neurohormone signal transduction for dual regulation of metabolism and gene expression in insects: hypertrehalosemic hormone as a model. Arch. Insect Biochem. Physiol. 33, 283–301. Kidd, G. and Devorak, J. (1994). Trehalose is a sweet target for agbiotech. Bio/ technology 12, 1328–1329. Kimura, M. T., Awasaki, T., Ohtsu, T. and Shimada, K. (1992). Seasonal changes in glycogen and trehalose content in relation to winter survival of four temperate species of Drosophila. J. Insect Physiol. 38, 871–875. Kingan, T. G., Zitnan, D., Jaffe, H. and Beckage, N. E. (1997). Identification of neuropeptides in the midgut of parasitized insects: FLRFamides as candidate paracrines. Molec. Cell. Endocrinol. 133, 19–32. Kloppenburg, P. and Hildebrand, J. G. (1995). Neuromodulation by 5-hydroxytryptamine in the antennal lobe of the sphinx moth Manduca sexta. J. Exp. Biol. 198, 603–611. Klowden, M. J. (2002). Physiological Systems in Insects. pp. 183. New York: Academic Press. Knapp, P. R. (1979). Handbook of Analytical Derivatization Reactions. pp. 539–543. New York: John Wiley. Kono, Y., Takahashi, M., Matsushita, K., Nishina, M., Kameda, Y. and Hori, E. (1994). Inhibition of flight in Periplaneta americana (Linn.) by a trehalase inhibitor, validoxylamine A. J. Insect Physiol. 40, 455–461.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
271
Kono, Y., Takahashi, M., Mihara, M., Matsushita, K., Nishina, M. and Kameda, Y. (1997). Effect of a trehalose inhibitor, validoxylamine A, on oocyte development and ootheca formation in Periplaneta americana (Blattodea, Blattidae). Appl. Entomol. Zool 32, 293–301. Kovalova, H. and Strunecka, A. (1973). A study of the distribution of haemolymph and total haemolymph volume of some insects. Acta Entomol. Bohemoslov 70, 334–337. Kramer, K. J. (1980). Insulin-like and glucagon-like hormones in insects. In: Neurohormonal Techniques in Insects (ed. Miller, T. A.), pp. 116–136. New York: Springer. Kramer, K. J. (1985). Vertebrate hormones in insects. In: Comprehensive Insect Biochemistry, Physiology and Pharmacology (eds. Kerkut, G. A., and Gilbert, L. I.), pp. 511–536. Kramer, K. J., Childs, C. N., Speirs, R. D. and Jacobs, R. M. (1982). Purification of insulin-like peptides from insect hemolymph and royal jelly. Insect Biochem. 12, 91–98. Kramer, K. J., Speirs, R. D. and Childs, C. N. (1978). A method for separation of trehalose from insect hemolymph. Anal. Biochem. 86, 692–696. Kramer, K. J., Tager, H. S. and Childs, C. N. (1980). Insulin-like and glucagon-like peptides in insect hemolymph. Insect Biochem. 10, 179–182. Kukal, O., Duman, J. G. and Serianni, S. (1989). Cold-induced mitochondrial degradation and cryoprotectant synthesis in freeze-tolerant arctic caterpillars. J. Comp. Physiol. 158B, 661–671. Kukal, O., Serianni, A. S. and Duman, J. G. (1988). Glycerol metabolism in a freezetolerant arctic insect: an in vivo 13C NMR study. J. Comp. Physiol. 158B, 175–183. Kulansky Poltilove, R. M., Jacobs, A. R., Haft, C. R., Xu, P. and Taylor, S. I. (2001). Characterization of Drosophila insulin receptor substrate. J. Biol. Chem. 275, 23346–23354. Kurahashi, H., Kawai, S. and Shudo, C. (1991). Seasonal migration of Japanese blow flies, Aldrichina grahami (Aldrich) and Calliphora nigribarbis Vollenhoven, observed by a mark and capture method on Hachijo Island, Tokyo. Jap. J. Sanit. Zool. 42, 57–59. La Course, W. R. (1997). Pulsed Electrochemical Detection in High Performance Liquid Chromatography. New York: John Wiley. Labat-Robert, J., Baumann, R. Ch., Bar-Guilloux, E. and Robic, D. (1978). Comparative specificities of trehalases from various species. Comp. Biochem. Physiol. 61B, 111–114. Landau, B. R. (1985). Use of radioisotopes in elucidating the nature of and quantitating the pentose pathway. In: The Pentose Phosphate Pathway (ed. Wood, T.), pp. 153–178. New York: Academic Press. Lee, J.-H., Tsuji, M., Nakamura, M., Nishimoto, M., Okuyama, M., Mori, H., Kimura, A., Matsui, H. and Chiba, S. (2001). Purification and identification of the essential ionizable groups of honeybee, Apis mellifera L., trehalase. Biosci. Biotechnol. Biochem. 65, 2657–2665. Lee, R. E. (1991). Principles of insect low temperature tolerance. In: Insects at Low Temperature (eds. Lee, R. E., and Dennlinger, D. L.), pp. 17–46. London: Chapman and Hall. Lee, R. E., Zachariassen, K. E. and Baust, J. G. (1981). Effect of cryoprotectants on the activity of hemolymph nucleating agents in physical solution. Cryobiology 18, 511–514. Lee, R. M. (1961). The variation of blood volume with age in the desert locust (Schistocerca gregaria Forsk.). J. Insect Physiol. 6, 36–51.
272
S. N. THOMPSON
Lee, Y.-H. and Keeley, L. L. (1994). Intracellular transduction of trehalose synthesis by hypertrehalosemic hormone in the fat body of the tropical cockroach, Blaberus discoidalis. Insect Biochem. Molec. Biol. 24, 473–480. Leevers, S. J. (2001). Growth control: invertebrate insulin surprises!. Curr. Biol. 11, R209–R212. Leta, M. A., Gilbert, C. and Morse, R. A. (1996). Levels of hemolymph sugars and body glycogen of honeybees (Apis mellifera L.) from colonies preparing to swarm. J. Insect Physiol. 42, 239–245. Levine, H. and Slade, L. (1992). Another view of trehalose for drying and stabilizing biological materials. Biopharm. 5, 36–40. Lewis, D. K., Bradfield, J. Y. and Keeley, L. L. (1998). Feeding effects on gene expression of the hypertrehalosemic hormone in the cockroach, Blaberus discoidalis. J. Insect Physiol. 44, 967–972. Lewis, J. G., Learmonth, R. P., Attfield, P. V. and Watson, K. (1997). Stress cotolerance and trehalose content in baking strains of Saccharomyces cerevisiae. J. Indus. Microbiol. Biotechnol. 18, 30–36. Liu, Q., Schmidt, R. K., Teo, B., Karplus, P. A. and Brady, J. W. (1997). Molecular dynamics studies of the hydration of -trehalose. J. Amer. Chem. Soc. 119, 7851–7862. Loomis, S. H. (1991). Comparative invertebrate cold hardiness. In: Insects at Low temperature (eds. Lee, R. E., and Denlinger, D. L.), pp. 301–317. New York: Chapman and Hall. Lum, P. Y. and Chino, H. (1990). Trehalose, the insect blood sugar, inhibits loading of diacylglycerol by lipophorin from the fat body in locusts. Biochem. Biophys. Res. Commun. 172, 588–594. Machin, J. (1981). Water compartmentalization in insects. J. Exp. Zool. 215, 327–333. Maddrell, S. H. P., Herman, W. S., Mooney, R. L. and Overton, J. A. (1991). 5-hydroxytryptamine: a second diuretic hormone in Rhodnius prolixus. J. Exp. Biol. 156, 557–566. Mangum, C. and Towle, D. (1977). Physiological adaptation to unstable environments. Amer. Sci. 65, 67–75. Mangum, C. P., Silverthorn, S. U., Harris, J. L., Towle, D. W. and Krall, A. R. (1976). The relationship between blood pH, ammonia excretion and adaptation to low salinity in the blue crab Callinectes sapidus. J. Exp. Zool. 195, 129–136. Maquenne, L. (1891). Recherches sur la trehalose. Compt. Rend. Hebd. Seanc. Acad. Sci 112, 947. Mariani, P., Rustichelli, F., Saturni, L. and Cordone, L. (1999). Stabilization of the monolein Pn3m cubic structure on trehalose glasses. Eur. Biophys. J. 28, 294–301. Masumura, M., Satake, S., Saegusa, H. and Mizoguchi, A. (2000). Glucose stimulates the release of bombyxin, an insulin-related peptide of the silkworm Bombyx mori. Gen. Comp. Endocrinol. 118, 393–399. McDougall, G. E. and Steele, J. E. (1988). Free fatty acids as a source of energy for trehalose synthesis in the fat body of the American cockroach (Periplaneta americana). Insect Biochem. 18, 591–597. Mellanby, K. (1939). The functions of insect blood. Biol. Rev. 14, 243–260. Mentel, T., Duch, C., Stypa, H., Wegener, G., Mu¨ller, U. and Pfu¨ger, H.-J. (2003). Central modulatory neurons control fuel selection in flight muscle of migratory locust. J. Neurosci. 23, 1109–1113. Meryman, H. T. (1971). Osmotic stress as a mechanism of freezing injury. Cryobiology 8, 489–500. Meryman, H. T. (1974). Freezing injury and its prevention in living cells. Ann. Rev. Biophys. Bioeng. 3, 341–363.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
273
Meyer zu Duttingdorf, H.-D., Bachmann, B., Buchholz, M. and Leuchtenberger, W. (1997). Determination of trehalose by flow injection analysis using immobilized trehalase. Anal. Biochem. 253, 8–12. Meyer-Fernandez, J. R., Arrese, E. L. and Wells, M. A. (2000). Allosteric effectors and trehalose protect larval Manduca sexta fat body glycogen phosphorylase B against thermal denaturation. Insect Biochem. Molec. Biol. 30, 473–478. Mikkat, S., Effmert, U. and Hagemann, M. (1997). Uptake and use of the osmoprotective compounds trehalose, glucosylglycerol, and sucrose by the cyanobacterium Synechocystis sp. PCC6803. Arch. Microbiol. 167, 112–118. Miller, L. K. (1982). Cold-hardiness strategies of some adult and immature insects overwintering in interior Alaska. Comp. Biochem. Physiol. 73A, 595–604. Miller, T. A. (1985). Structure and physiology of the circulatory system. In: Comparative Insect Physiology, biochemistry and Pharmacology, Vol. 3 (eds. Kerkut, G. A., and Gilbert, L. I.), pp. 289–353. New York: Pergamon Press. Miller, T. A. (1997). Control of circulation in insects. Gen. Pharmacol. 29, 23–38. Moelwyn-Hughes, E. A. (1929). Kinetics of the hydrolysis of certain glucosides. III -methylglucoside, cellobiose, melibiose and turanose. Trans. Faraday Soc. 25, 503–520. Moelwyn-Hughes, E. A. (1930). Note on the variation with temperature of the relative rates of hydrolysis of glucosides. J. Gen. Physiol. 13, 317–334. Moreau, R., Gourdoux, L. and Cava-Ruart, M. (1980). Neuroendocrine control possibility of the glucose degradation pathways in non diapausic Pieris brassicae L. (Lepidoptera). Comp. Biochem. Physiol. 66B, 589–592. Mori, H. (1979). Embryonic hemocytes: origin and development. In: Insect Hemocytes (ed. Gupta, A. P.), pp. 3–27. Cambridge: Cambridge University Press. Morita, H. and Shiraishi, A. (1968). Stimulation of the labellar sugar receptor of the fleshfly by mono- and diisaccharides. J. Gen. Physiol. 52, 559–583. Mouradian, R., Womersley, C., Crowe, L. M. and Crowe, J. H. (1985). Degradation of functional integrity during long-term storage of a freeze-dried biological membrane. Cryobiology 22, 119–127. Mtioui, A., Bahjou, A., Gourdoux, L. and Moreau, R. (1994). Comparison of hormonal effects on glucose metabolism in Tenebrio molitor larval fat body, muscle and brain tissues. Comp. Biochem. Physiol. 108A, 97–105. Mtioui, A., Gourdoux, L., Fournier, B. and Moreau, R. (1993a). Effects of intestinal insulin-like peptide on glucose catabolism in mealworm larval fat body in vitro: dependence on extracellular Ca2 þ for its stimulatory action. Arch. Insect Biochem. Physiol. 24, 113–128. Mtioui, A., Gourdoux, L., Moreau, R. and Fournier, B. (1996). Relationships between adenylyl cyclase activity, glucose metabolism, and both intestinal insulin-like peptide and adipokinetic hormone in mealworm larval fat body. Arch. Insect Biochem. Physiol. 32, 209–223. Mtioui, A., Moreau, R., Bahjou, A. and Gourdoux, L. (1993b). Regulation of mealworm fat body glucose metabolism in vitro: effects of natural and synthetic catabolic neurohormones. J. Exp. Zool. 267, 493–500. Muise, A. M. and Storey, K. B. (1997). Reversible phosphorylation of fructose 1,6bisphosphatase mediates enzyme role in glycerol metabolism in the freeze-avoiding gall moth Epiblema scudderiana. Insect Biochem. Molec. Biol. 27, 617–623. Muise, A. M. and Storey, K. B. (1999). Regulation of glycogen synthetase in a freezeavoiding insect: role in crypoprotectant glycerol metabolism. Cryo-Lett. 20, 223–228. Muise, A. M. and Storey, K. B. (2001). Regulation of hexokinase in a freeze avoiding insect: role in the winter production of glycerol. Arch. Insect Biochem. Physiol. 47, 29–34.
274
S. N. THOMPSON
Muller, J., Boller, T. and Wiemken, A. (1995). Trehalose and trehalase in plants: recent developments. Plant Sc. 112, 1–9. Mullins, D. E. (1985). Chemistry and physiology of the hemolymph. In: Comparative Insect Physiology, Biochemistry and Pharmacology, Vol. 3 (eds. Kerkut, G. A., and Gilbert, L. I.), pp. 355–400. New York: Pergamon Press. Murphy, T. A. and Wyatt, G. R. (1965). The enzymes of glycogen and trehalose synthesis in silk moth fat body. J. Biol. Chem. 240, 1500–1508. Murray, D. B., Hayashida, Y. and Nishimura, K. (1997). Trehalose analysis using ion exchange HPLC coupled with electrochemical detection. Biotech. Tech. 11, 269–270. Nagasawa, H., Kataoka, H., Isogai, A., Tamura, S., Suzuki, A., Mizoguchi, A., Fujiwara, Y., Suzuki, A., Takahashi, S. Y., Ishizaki, H. (1986). Amino acid sequence of a prothoracicotropic hormone of the silkworm Bombyx mori. Proc. Nat. Acad. Sci. 83, 5840–5843. Nagasawa, H., Kataoka, H. and Suzuki, A. (1990). Chemistry of Bombyx prothoracicotropic hormone and bombyxin. In: Molting and Metamorphosis (eds. Ohnishi, E., and Ishizake, H.), pp. 33–48. Berlin: Springler-Verlag. Nagata, K., Hatanaka, H., Kohda, D., Kataoka, H., Nagasawa, H., Isogai, A., Ishizaki, H., Suzuki, A. and Inagaki, F. (1995a). Three-dimensional solution structure of bombyxin-II an insulin-like peptide of the silkmoth Bombyx mori – structural comparison with insulin and relaxin. J. Molec. Biol. 253, 749–758. Nagata, K., Hatanaka, H., Kohda, D., Kataoka, H., Nagasawa, H., Isogai, A., Ishizaki, H., Suzuki, A. and Inagaki, F. (1995b). Identification of the receptorrecognition surface of bombyxin-II, an insulin-like peptide of the silkmoth, Bombyx mori: critical importance of the B-chain central part. J. Molec. Biol. 759–770. Nathanson, J. A. (1993). Identification of octopaminergic agonists with selectivity for octopamine receptor subtypes. J. Pharmacol. Exp. Ther. 265, 509–515. Nation, J. L. (2002). Insect Physiology and Biochemistry. Boca Raton: CRC Press. Nayar, J. K. and Sauerman, D. M. (1971). Physiological effects of carbohydrates on survival, metabolism, and flight potential of female Aedes taeniorhynchus. J. Insect Physiol. 17, 2221–2233. Nijhout, H. F. (1994). Insect hormones. Princeton: Princeton University Press. Nomura, M. and Ishikawa, Y. (2001). Dynamic changes in cold hardiness, hightemperature tolerance and trehalose content in the onion maggot, Delia antiqua (Diptera: Anthomyiidae), associated with the summer and winter diapause. Appl. Entomol. Zool. 36, 443–449. Normann, T. C. (1975). Neurosecretory cells in insect brain and production of hypoglycaemic hormone. Nature 254, 259–261. O’Brien, J. (1996). Stability of trehalose, sucrose and glucose to nonenzymatic browning in model systems. J. Food Sci. 61, 679–682. Oda, Y., Uejuma, M., Iwami, M. and Sakurai, S. (2000). Involvement of adipokinetic hormone in the homeostatic control of hemolymph trehalose concentration in larvae of Bombyx mori. Arch. Insect Biochem. Physiol. 45, 156–165. Okajima, A., Kumagai, K. and Watanabe, N. (1989). The involvement of interoceptive chemosensory activity in the nervous regulation of the prothoracic gland in a moth, Mamestra brassicae. Zool. Sc. 6, 859–866. Orchard, I. (1987). Adipokinetic hormones – an update. J. Insect Physiol. 33, 451–463. Orchard, I., Lange, A. B. and Bendena, W. G. (2001). FMRFamide-related peptides: a multifunctional family of structurally related neuropeptides in insects. Adv. Insect Physiol. 28, 267–329.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
275
Orchard, I., Loughton, B. G. and Webb, R. A. (1981). Octopamine and short-term hyperlipaemia in the locust. Gen. Comp. Endocrinol. 45, 175–180. Orchard, I., Ramirez, J.-M. and Lange, A. B. (1993). A multifunctional role for octopamine in locust flight. Ann. Rev. Entomol. 38, 227–249. Panek, A. D. (1995). Trehalose metabolism – new horizons in technological applications. Braz. J. Med. Biol. Res. 28, 169–181. Panek, A. D. and Panek, A. C. (1990). Metabolism and thermotolerance function of trehalose in Saccharomyces: a current perspective. J. Biotech. 14, 229–238. Park, J. H. and Keeley, L. L. (1996a). Calcium-dependent action of hypertrehalosemic hormone on activation of glycogen phosphorylase in cockroach fat body. Molec. Cell. Endocrinol. 116, 199–205. Park, J. H. and Keeley, J. H. (1996b). The effect of biogenic amines and their analogs on carbohydrate metabolism in the fat body of the cockroach Blaberus discoidalis. Gen. Comp. Endocrinol. 110, 88–95. Pass, G. (2000). Accessory pulsatile organs: evolutionary innovations in insects. Annu. Rev. Entomol. 45, 495–518. Passier, P. C. C. M., Vullings, H. G. B., Diederen, J. H. B. and van der Horst, D. J. (1997). Trehalose inhibits the release of adipokinetic hormones from the corpus cardiacum in the African migratory locust, Locusta migratoria, at the level of the adipokinetic cells. J. Endocrinol. 153, 299–305. Pauli, D., Arrigo, A.-P. and Tissieres, A. (1992). Heat shock response in Drosophila. Experimentia 48, 623–629. Pfister, T. D. and Storey, K. B. (2002). Purification and characterization of protein phosphatase-1 from two cold-hardy goldenrod gall insects. Arch. Biochem. Physiol. 49, 56–64. Phalaraksh, C., Lenz, E. M., Lindon, J. C., Nicholson, J. K., Farrant, R. D., Reynolds, S. E., Wilson, I. D., Osborn, D. and Weeks, J. M. (1999). NMR spectroscopic studies on the haemolymph of the tobacco hornworm, Manduca sexta: assignment of 1H and 13C NMR spectra. Insect Biochem. Molec. Biol. 29, 795–805. Pierce, W. D. (1915). The uses of certain weevils and weevil products in food and medicine. Proc. Entomol. Soc. Wash. 17, 151–154. Pilkis, S. J. and Claus, H. T. (1995). 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase – a metabolic signaling enzyme. Ann. Rev. Biochem. 64, 799–835. Pilkis, S. J. and Claus, T. H. (1991). Hepatic gluconeogenesis/glycolysis: regulation and structure/function relationships of substrate cycle enzymes. Ann. Rev. Nutr. 11, 465–515. Portmann, M.-O. and Birch, G. (1995). Sweet taste and solution properties of , trehalose. J. Sci. Food Agric. 69, 275–281. Prestrelski, S. J., Arakawa, T. and Carpenter, J. F. (1993). Separation of freezing and drying-induced denaturation of lypophilized proteins using stress-specific stabilization. Structural studies using infrared spectroscopy. Arch. Biochem. Biophys. 303, 456–464. Quinn, P. J. (1985). A lipid-phase separation model of low-temperature damage to biological membranes. Cryobiology 22, 128–146. Raubenheimer, D. and Simpson, S. J. (1990). The effects of simultaneous variation in protein, digestible carbohydrate and tannic acid on the feeding behaviour of larval Locusta migratoria (L.). and Schistocerca gregaria (Forskal.). I. Short-term studies. Physiol. Entomol. 15, 219–233. Raubenheimer, D. and Simpson, S. J. (1993). The geometry of compensatory feeding in the locust. Anim. Behav. 45, 953–964. Raubenheimer, D. and Simpson, S. J. (1999). Integrating nutrition: a geometrical approach. Entomol. Exp. Appl. 91, 67–82.
276
S. N. THOMPSON
Reed, W. D. and Sacktor, B. (1971). Localization of trehalase in flight muscle of the blowfly, Phormia regina. Archs. Biochem. Biophys. 145, 392–401. Reinecke, J. P. (1985). Nutrition: artificial diets. In: Comparative Insect Physiology, Biochemistry and Pharmacology, Vol. 4 (eds. Kerkut, G. A., and Gilbert, L. I.), pp. 391–419. New York: Pergamon Press. Reinecke, T. P., Cook, B. J. and Adams, T. S. (1973). Larval hindgut of Manduca sexta (L.) (Lepidoptera: Sphingidae). J. Insect Morphol. Embryol. 2, 277–290. Rheuben, M. B. (1995). Specific associations of neurosecretory or neuromodulatory axons with insect skeletal muscle. Amer. Zool. 35, 566–577. Ring, R. A. and Danks, H. V. (1998). The role of trehalose in cold-hardiness and desiccation. Cryo-Letters 19, 275–282. Roeder, T. (1999). Octopamine in invertebrates. Prog. Neurobiol. 59, 533–561. Rosser, B. (1991). Trehalose, a new approach to premium dried foods. TFST 2, 166–169. Rowell, C. H. F. and Simpson, S. J. (1992). A peripheral input of thoracic origin inhibits chewing movements in the larvae of Manduca sexta. J. Insect Physiol. 38, 475–483. Rulifson, E. J., Kim, S. K. and Nusse, R. (2002). Ablation of insulin-producing neurons in flies: growth and diabetic phenotypes. Science 296, 1118–1120. Sacktor, B. (1975). Biochemistry of insect flight. Part 1 Utilization of fuels by muscle. In: Insect Biochemistry and Function (eds. Candy, D. J., and Kilby, B. A.), pp. 1–88. New York: Chapman and Hall. Sacktor, B. (1976). Biochemical adaptations for flight in the insect. Biochem. Soc. Symp. 41, 111–131. Sacktor, B. and Wormser-Shavit, E. (1966). Regulation of metabolism in working muscle in vivo. I. concentration of some glycolytic, tricarboxylic acid cycle, and amino acid intermediates in insect flight muscle during flight. J. Biol. Chem. 241, 624–631. Saito, S. (1960). Trehalase of the silkworm, Bombyx mori: purification and properties of the enzyme. J. Biochem. 48, 101–109. Sakurai, M., Murata, M., Inoue, Y., Hino, A. and Kobayashi, S. (1997). Moleculardynamics study of aqueous solution of trehalose and maltose: implication for the biological function of trehalose. Bull. Chem. Soc. Jap. 70, 847–858. Salam, S. E. A., Moto, K., Sakurai, S. and Iwami, M. (2001). Transcription element responsible for the brain cell-specific expression of the bombyxin gene that encodes an insect insulin-related peptide. Zool. Sci. 18, 543–549. Sanderson, P. W., Lis, L. J., Quinn, P. J. and Williams, W. P. (1991). The Hofmeister effect in relation to membrane lipid phase stability. Biochim. Biophys. Acta 1067, 43–50. Sastry, G. M. and Agmon, N. (1997). Trehalose prevents myoglobin collapse and preserves its internal mobility. Biochemistry 36, 7097–7108. Satake, S., Kawabe, Y. and Mizoguchi, A. (2000). Carbohydrate metabolism during starvation in the silkworm Bombyx mori. Arch. Insect Biochem. Physiol. 44, 90–98. Satake, S., Masumura, M., Ishizaki, H., Nagata, K., Kataoka, H., Suzuki, A. and Mizoguchi, A. (1997). Bombyxin, an insulin-related peptide of insects, reduces the major storage carbohydrates in the silkworm Bombyx mori. Comp. Biochem. Physiol. 118B, 349–357. Satake, S., Nagata, K., Kataoka, H. and Mizoguchi, A. (1999). Bombyxin secretion in the adult silkmoth Bombyx mori: sex-specificity and its correlation with metabolism. J. Insect Physiol. 45, 939–945. Sato, K., Komoto, M., Sato, T., Enei, H., Michihiro, K. and Yaginuma, T. (1997). Baculovirus-mediated expression of a gene for trehalase of the mealworm beetle,
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
277
Tenebrio molitor, in insect cells, SF-9, and larvae of the cabbage armyworm, Mamestra brassicae. Insect Biochem. Molec. Biol. 27, 1007–1016. Schiff, N. M., Waldbauer, G. P. and Friedman, S. (1989). Dietary self-selection by Heliothis zea larvae: roles of metabolic feedback and chemosensory stimuli. Entomol. Exp. Appl. 52, 261–270. Schooneveld, H., Romberg-Privee, H. M. and Veenstra, J. A. (1987). Phylogenetic differentiation of glandular cells in corpora cardiaca as studied immunocytochemically with region-specific anisera to adipokinetic hormone. J. Insect Physiol. 33, 167–176. Schulze, U., Larsen, M. E. and Villadsen, J. (1995). Determination of intracellular trehalose and glycogen in Saccharomyces cerevisiae. Anal. Biochem. 228, 142–149. Sehnal, F. and Zitnan, D. (1990). Endocrines of insect gut. In: Progress in Comparative Endocrinology (eds. Epple, A., Scanes, C. G., and Stetson, M. H.), pp. 510–515. New York: Wiley-Liss. Sevala, V. L. and Steele, J. E. (1989). Absence of a stimulatory effect of the corpus cardiacum on gluconeogenesis in cockroach (Periplaneta americana) fat body. J. Insect Physiol. 35, 1031–1036. Sevala, V. M. and Loughton, B. G. (1992). Insulin-like peptides during oogenesis and embryogenesis in Locusta migratoria. Invert. Repro. Develop. 21, 187–192. Sevala, V. M., Sevala, V. L. and Loughton, B. G. (1993). Insulin-like molecules in the beetle Tenebrio molitor. Cell Tiss. Res. 273, 71–77. Shimada, S. and Yamashita, O. (1979). Trehalose absorption related with trehalase in developing ovaries of the silkworm, Bombyx mori. J. Comp. Physiol. 131, 333–339. Shiraishi, A. and Yano, T. (1984). Neuronal control of the feeding behavior in the blow fly. In: Animal Behavior: Neurophysical and Ethological Approaches (eds. Aoki, K., Ishii, S., and Morita, H.), pp. 83–93. Berlin: Springer-Verlag. Shor-Posner, G., Grinker, J. A., Marinescu, C., Brown, O. and Leibowitz, S. F. (1986). Hypothalamic serotonin in the control of meal patterns and macronutrient selection. Brain Res. Bull. 17, 663–671. Siegert, K. J. (1987). Carbohydrate metabolism in starved fifth instar larvae of Manduca sexta. Arch. Insect Biochem. Physiol. 4, 151–160. Siegert, K. J. and Mordue, W. (1992). Regulation of fat body glycogen phosphorylase activity during refeeding in Manduca sexta larvae. Arch. Insect Biochem. Physiol. 19, 225–235. Siegert, K. J., Krippeit, P. and Ziegler, R. (1982). Regulation of fat body glycogen phosphorylase during starvation in larvae of Manduca sexta (Lepidoptera: Sphingidae). Acta Endocrinol. 246S, 32–33. Simmonds, M. S. J., Simpson, S. J. and Blaney, W. M. (1992). Dietary selection behaviour in Spodoptera littoralis: the effects of conditioning diet and conditioning period on neural responsiveness and selection behaviour. J. Exp. Biol. 162, 73–90. Simola, M., Hanninen, A.-L., Stranius, S.-M. and Makarow, M. (2000). Trehalose is required for conformational repair of heat-denatured proteins in the yeast endoplasmic reticulum but not for maintenance of membrane traffic functions after severe heat stress. Molec. Microbiol. 37, 42–53. Simpson, C. L., Chyb, S. and Simpson, S. J. (1990a). Changes in chemoreceptor sensitivity in relation to dietary selection by adult Locusta migratoria. Entomol. Exp. Appl. 56, 259–268. Simpson, S. J. (1995). Regulation of a meal. In: Regulatory Mechanisms in Insect Feeding (eds. Chapman, R. F., and de Boer, G.), Chapter 5, pp. 137–156. New York: Chapman and Hall.
278
S. N. THOMPSON
Simpson, S. J. and Raubenheimer, D. (1993a). A multi-level analysis of feeding behaviour: the geometry of nutritional decisions. Phil. Trans. Roy. Soc. 342, 381–402. Simpson, S. J. and Raubenheimer, D. (1993b). The central role of the haemolymph in the regulation of nutrient intake in insects. Physiol. Entomol. 18, 395–403. Simpson, S. J. and Raubenheimer, D. (1996). Feeding behaviour, sensory physiology and nutrient feedback: a unifying model. Entomol. Exp. Appl. 80, 55–64. Simpson, S. J. and Simpson, C. L. (1992). Mechanisms controlling modulation by amino acids of gustatory responsiveness in the locust. J. Exp. Biol. 168, 269–287. Simpson, S. J., James, S., Simmonds, M. S. J. and Blaney, W. M. (1991). Variation in chemosensitivity and the control of dietary selection behaviour in the locust. Appetite 17, 141–154. Simpson, S. J., Raubenheimer, D. and Chambers, P. G. (1995). The mechanisms of nutritional homeostasis. In: Regulatory Mechanisms in Insect Feeding (eds. Chapman, R. F., and de Boer, G.), Chapter 9, pp. 251–278. New York: Chapman and Hall. Simpson, S. J., Simmonds, M. S. J. and Blaney, W. M. (1988). A comparison of dietary selection behaviour in larval Locusta migratoria and Spodoptera littoralis. Physiol. Entomol. 13, 225–238. Simpson, S. J., Simmonds, M. S. J., Blaney, W. M. and Jones, J. P. (1990b). Compensatory dietary selection occurs in larval Locusta migratoria but not Spodoptera littoralis after a single deficient meal during ad libitum feeding. Physiol. Entomol. 15, 235–242. Singer, M. A. and Lindquist, S. (1998a). Thermotolerance in Saccharomyces cerevisiae: the yin and yang of trehalose. TIBTech. 16, 460–468. Singer, M. A. and Lindquist, S. (1998b). Multiple effects of trehalose on protein folding in vitro and in vivo. Molec. Cell 1, 639–648. Singh, P. (1977). Artificial diets for insects, mites, and spiders. New York: Plenum Press. Slansky, F. and Rodriguez, J. G. (1987). Nutritional ecology of insects, mites, spiders, and related invertebrates: an overview. In: Nutritional Ecology of Insects, Mites, Spiders, and Related Invertebrates (eds. Slansky, F., and Rodriguez, J. G.), pp. 1–69. New York: John Wiley and Sons. Sola-Penna, M. and Meyer-Fernandes, J. R. (1998). Stabilization against thermal inactivation promoted by sugars on enzyme structure and function: Why is trehalose more effective than other sugars? Arch. Biochem. Biophys. 360, 10–14. Somme, L. (1982). Supercooling and winter survival in terrestrial arthropods. Comp. Biochem. Physiol. A 73, 519–543. Sonobe, H. A., Matsumoto, A., Fukuzaki, Y. and Fujiwara, S. (1979). Carbohydrate metabolism and restricted oxygen supply in the eggs of the silkworm, Bombyx mori. J. Insect Physiol. 25, 381–388. Sowa, S. M., Lu, K.-H., Park, J. H. and Keeley, L. L. (1996). Physiological effectors of hyperglycemic neurohormone biosynthesis in an insect. Molec. Cell. Endocrinol. 123, 970105. Steele, J. E. (1963). The site of action of insect hyperglycaemic hormone. Gen. Comp. Endocrinol. 3, 46–52. Steele, J. E. (1980). Hormonal modulation of carbohydrate and lipid metabolism in the fat body. In: Insect Biology in the Future (eds. Locke, M., and Smith, D. S.), pp. 253–271. New York: Academic Press. Steele, J. E. (1981). The role of carbohydrate metabolism in physiological function. In: Energy Metabolism in Insects (ed. Downer, R. G. H.), pp. 101–133. New York: Plenum. Steele, J. E. (1982). Glycogen phosphorylase in insects. Insect Biochem. 12, 131–147.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
279
Steele, J. E. and Hall, S. (1985). Trehalose synthesis and glycogenolysis as sites of action for the corpus cardiacum in Periplaneta americana. Insect Biochem. 15, 529–536. Steele, J. E. and Paul, T. (1985). Corpus cardiacum stimulated trehalose efflux from cockroach (Periplaneta americana). fat body: control by calcium. Can. J. Zool. 63, 63–66. Steele, J. E., Garcha, K. and Sun, D. (2001). Inositol triphosphate mediates the action of hypertrehalosemic hormone on fat body of the American cockroach, Periplaneta americana. Comp. Biochem. Physiol. 130B, 537–545. Storey, J. M. and Storey, K. B. (1983). Regulation of cryoprotectant metabolism in the overwintering gall fly larvae, Eurosta solidaginis: temperature control of glycerol and sorbitol levels. J. Comp. Physiol. 149, 495–502. Storey, K. B. (1997). Organic solutes in freezing tolerance. Comp. Biochem. Physiol. 117A, 319–326. Storey, K. B. and Bailey, E. (1978a). Intracellular distribution of enzymes associated with lipogenesis and gluconeogenesis in fat body of the adult cockroach, Periplaneta. Insect Biochem. 8, 125–131. Storey, K. B. and Bailey, E. (1978b). The intracellular distribution of enzymes of carbohydrate degradation in the fat body of the adult male cockroach. Insect biochem. 8, 73–79. Storey, K. B. and Storey, J. M. (1988). Freeze tolerance in animals. Physiol. Rev. 68, 27–84. Storey, K. B. and Storey, J. M. (1991). Biochemistry of cryoprotectants. In: Insects at Low Temperature (eds. Lee, R. E., and Dennlinger, D. L.), pp. 64–93. London: Chapman and Hall. Storey, K. B., Baust, J. G. and Storey, J. M. (1981). Intermediary metabolism during low temperature acclimation in the overwintering gall fly larvae, Eurosta solidaginis. J. Comp. Physiol. 144, 191–199. Su, H. Z., Sato, Y. and Yamashita, O. (1993). Purification, cDNA cloning and northern blot analysis of trehalase of pupal midgut of the silkworm, Bombyx mori. Biochim. Biophys. Acta 1173, 217–224. Su, H.-Z., Ikeda, M., Sato, Y., Saito, H., Imai, K., Isobe, M. and Yamashita, O. (1994). Molecular characterization of ovary trehalase of the silk worm, Bombyx mori, and its transcriptional activation by diapause hormone. Biochim. Biophys. Acta 1218, 366–374. Suarez, R. K. (2000). Energy metabolism during insect flight: biochemical design and physiological performance. Physiol. Biochem. Zool. 73, 765–771. Suarez, R. K., Lighton, J. R. B., Joos, B., Roberts, S. P. and Harrison, J. F. (1996). Energy metabolism, enzymatic flux capacities and metabolic flux rates in flying honeybees. Proc. Natl. Acad. Sci. 93, 12616–12620. Sun, D., Garcha, K. and Steele, J. E. (2002). Stimulation of trehalose efflux from cockroach (Periplaneta americana) fat body by hypertreahlosemic hormone is dependent on protein kinase C and calmodulin. Arch. Insect Biochem. Physiol. 50, 41–51. Surholt, B., Greive, H., Baal, T. and Bertsch, A. (1990). Non-shivering thermogenesis in asynchronous flight muscles of bumblebees? Comparative studies on males of Bombus terrestris, Xylocopa sulcatipes and Acherontia atropos. Comp. Biochem. Physiol. 97A, 439–499. Sussich, F., Princivalle, F. and Cesaro, A. (1999). The interplay of the rate of water removal in the dehydration of , -trehalose. Carb. Res. 322, 113–119. Sussich, F., Skopec, C., Brady, J. and Cesaro, A. (2001). Reversible dehydration of trehalose and anhydrobiosis; from solution state to an exotic crystal? Carb. Res. 334, 165–176.
280
S. N. THOMPSON
Sutcliffe, D. W. (1963). The chemical composition of haemolymph in insects and some other arthropods, in relation to their phylogeny. Comp. Biochem. Physiol. 9, 121–135. Sutherland, E. W. (1972). Studies on the mechanism of hormone action. Science 177, 401–408. Sweeley, C. C., Bentely, R., Makita, M. and Wells, W. W. (1963). Gas-liquid chromatography of trimethylsilyl derivatives of sugars and related substances. J. Amer. Chem. Soc. 85, 2497–2507. Taga, T., Senma, M. and Osaki, K. (1972). The crystal and molecular structure of trehalose dihydrate. Acta Cryst. B28, 3258–3263. Tager, H. S. and Kramer, K. J. (1980). Insect glucagon-like peptides: evidence for high molecular weight form in midgut from Manduca sexta (L.). Insect Biochem. 10, 617–619. Taiz, L. and Zeiger, E. (1998). Plant Physiology. pp. 259. Sunderland: Sinauer Associates. Takahashi, M., Kono, Y., Kurahashi, H., Matsushita, K., Nishina, M. and Kameda, Y. (1995). Effect of a trehalase inhibitor, validoxylamine A, on three species of flies. Appl. Entomol. Zool. 30, 231–239. Takahashi, S. Y., Higashi, S., Minoshima, S., Ogiso, M. and Hanaoka, K. (1980). Trehalases from the Amercian cockroach, Periplaneta americana: multiple occurrence of the enzymes and partial purification of enzymes from male accessory glands. Int. J. Invert. Reprod. 2, 373–381. Takenaka, S., Kondo, T., Nazeri, S., Tamura, Y., Tokunaga, M., Tsuyama, S., Miyatake, K. and Nakano, Y. (1997). Accumulation of trehalose as a compatible solute under osmotic stress in Euglena gracilis Z. J. Euk. Microbiol. 44, 609–613. Takiguchi, M., Niimi, T., Su, H.-Z. and Yaginuma, T. (1992). Trehalase from the accessory gland of an insect Tenebrio molitor. cDNA sequencing and developmental profile of the gene expression. Biochem. J. 288, 19–22. Talbot, B. G. and Huber, R. E. (1975). Partial purification, stabilization and characterization of the adult honey bee midgut trehalase and a new trehalose specific disc gel stain method. Insect Biochem. 5, 337–347. Taylor, M. J. (1987). Physio-chemical principles in low temperature biology. In: The Effects of Low Temperatures on Biological Systems (eds. Grout, B. W. W., and Morris, G. J.), pp. 3–71. London: Edward Arnold. Telfer, W. H. and Kunkel, J. G. (1991). The function and evolution of insect storage hexamers. Ann. Rev. Entomol. 36, 205–228. Tembhare, D. B. and Andrew, R. J. (1991). Homonal influence on the haemolymph trehalose concentration in the dragonfly Tramea virginia (Rambur) larva. Entomon 16, 165–166. Terra, W. R. and Ferreira, C. (1994). Insect digestive enzymes – properties, compartmentalization and function. Comp. Biochem. Physiol. 109B, 1–62. Thevelein, J. M. (1984). Regulation of trehalose mobilization in fungi. Microbiol. Rev. 48, 42–59. Thompson, S. N. (1985). Effects of AMP and fructose 2,6 bisphosphate on fructose 1,6 bisphosphatase activity in Trichoplusia ni parasitized by the insect parasite, Hyposoter exiguae. J. Parasitol. 71, 117–119. Thompson, S. N. (1990). NMR spectroscopy: its basis, biological application and use in studies of insect metabolism. Insect Biochem. 20, 223–237. Thompson, S. N. (1995). Gluconeogenesis and effect of nutritional status on TCA cycle activity in the insect Manduca sexta. Biochim. Biophy. Acta 1245, 376–384.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
281
Thompson, S. N. (1997a). Absence of short-term regulation over gluconeogenesis by glucose in the insect Manduca sexta L. Biochem. Biophys. Res. Commun. 237, 702–706. Thompson, S. N. (1997b). Glucogenesis in an insect, Manduca sexta L., estimated from the 13C isotopomer distribution in trehalose synthesized from [1,3-13C2]glycerol. Biochim. Biophys. Acta 1336, 110–116. Thompson, S. N. (1998). Long-term regulation of glucogenesis by dietary carbohydrate and relevance to blood sugar level in an insect Manduca sexta L. Int. J. Biochem. Cell Biol. 30, 987–999. Thompson, S. N. (1999). Blood sugar formation from dietary carbohydrate is facilitated by the pentose phosphate pathway in an insect Manduca sexta Linnaeus. Biochim. Biophys. Acta 1472, 565–575. Thompson, S. N. (2000). Pyruvate cycling and implications for regulation of gluconeogenesis in the insect, Manduca sexta. Biochem. Biophys. Res. Comm. 274, 787–793. Thompson, S. N. and Binder, B. F. (1984). Altered carbohydrate levels and gluconeogenic enzyme activity in Trichoplusia ni parasitized by the insect parasite, Hyposoter exiguae. J. Parasitol. 70, 644–651. Thompson, S. N. and Redak, R. A. (2000). Interactions of dietary protein and carbohydrate determine blood sugar level and regulate nutrient selection in the insect Manduca sexta L. Biochim. Biophys. Acta 1523, 91–102. Thompson, S. N., Borchardt, D. B. and Wang, L.-W. (2003). Dietary nutrient levels regulate protein and carbohydrate intake, gluconeogenic/glycolytic flux and blood trehalose level in the insect Manduca sexta L. J. Comp. Physiol. B. 173, 149–163. Thompson, S. N., Redak, R. A. and Borchardt, D. B. (2002). The glucogenic response of a parasitized insect Manduca sexta L. is partially mediated by differential nutrient intake. Biochim. Biophys. Acta 1571, 138–150. Thompson, S. N., Redak, R. A. and Wang, L.-W. (2001). Altered dietary nutrient intake maintains metabolic homeostasis in parasitized larvae of the insect Manduca sexta L. J. Exp. Biol. 204, 4065–4080. Timmins, W. A. and Reynolds, S. E. (1992). Physiological mechanisms underlying the control of meal size in Manduca sexta larvae. Physiol. Entomol. 17, 81–89. Timmins, W. A., Bellward, K., Stamp, A. J. and Reynolds, S. E. (1988). Food intake, conversion efficiency, and feeding behaviour of tobacco hornworm caterpillars given artificial diet of varying nutrient and water content. Physiol. Entomol. 13, 303–314. Treherne, J. E. (1967). Gut absorption. Ann. Rev. Entomol. 12, 43–58. Tsonev, L. I. and Tihova, M. G. (1994). The effect of the cryoprotective sugar trehalose on the phase behaviour of mixed dispersions of dioleoyl derivatives of phosphatidylethanolamine and phosphatidylcholine. Liq. Crys. 17, 717–728. Tsumuki, H. and Kanehisa, K. (1981). The fate of 14C-glycerol in the rice stem borer, Chilo suppressalis Walker (Lepidoptera: Pyralidae). Appl. Entomol. Zool. 16, 200–208. Tsumuki, H., Rojas, R. R., Storey, K. B. and Baust, J. G. (1987). The fate of [14C]glucose during cold-hardening in Eurosta solidaginis (Fitch). Insect Biochem. 17, 347–352. Tu, Z. and Hagedron, H. H. (1992). Purification and characterization of pyruvate carboxylase from the honeybee and some properties of related biotin-containing proteins in other insects. Arch. Insect Biochem. Physiol. 19, 53–66. Tunnacliffe, A., Garcia de Castro, A. and Manzanera, M. (2001). Anhydrobiotic engineering of bacterial and mammalian cells: is intracellular trehalose sufficient. Cryobiology 43, 124–132.
282
S. N. THOMPSON
Turunen, S. (1985). Absorption. In: Comparative Insect Physiology, Biochemistry and Pharmacology (eds. Kerkut, G. A., and Gilbert, L. I.), Vol. 4. pp. 241–277. New York: Pergamon Press. Turunen, S. (1993). Metabolic pathways in the midgut epithelium of Pieris brassicae during carbohydrate and lipid assimilation. Insect Biochem. Molec. Biol. 23, 681–689. Ueno, K., Ohta, M., Morita, H., Mikuni, Y., Nakajima, S., Yamamoto, K. and Isono, K. (2001). Trehalose sensitivity in Drosophila correlates with mutations in and expression of the gustatory receptor gene Gr5a. Curr. Biol. 11, 1451–1455. Uritani, M., Takai, M. and Yoshinaga, K. (1995). Protective effect of disaccharides on restriction endonucleases during drying under vacuum. J. Biochem. 117, 774–779. Vaandrager, S. H., Haller, T. B., van Marrewijk, W. J. A. and Beenakkers, A. M. Th. (1989). Fractionation and kinetic properties of trehalase from flight muscles and haemolymph of the locust, Locusta migratoria. Insect Biochem. 19, 89–94. van der Horst, D. J., van Doorn, J. M. and Beenakkers, A. M. Th. (1979). Effects of the adipokinetic hormone on the release and turnover of haemolymph diglycerides and on the formation of the diglyceride-transporting lipoprotein system during locust flight. Insect Biochem. 9, 627–635. van der Horst, D. J., van Doorn, J. M. and Beenakkers, A. M. T. (1978). Dynamics in the hemolymph trehalose pool during flight of the locust Locusta migratoria. Insect Biochem. 8, 413–416. van Handel, E. (1965). Microseparation of glycogen, sugars and lipids. Anal. Biochem. 11, 266–271. van Handel, E. (1978). Trehalose turnover and the coexistence of trehalose and active trehalase in cockroach haemolymph. J. Insect Physiol. 24, 151–153. van Laere, A. (1989). Trehalose, reserve and/or stress metabolite. Microbiol. Rev. 63, 201–210. van Marrewijk, W. J. A., van den Broek, A. T. M. and van der Horst, D. J. (1993). Adipokinetic hormone-induced influx of extracellular calcium is mediated through depletion of intracellular calcium stores. Cell. Signal. 5, 753–761. van Schaftingen, E. (1990). Role of fructose 2,6 bisphosphate in regulation of hepatic carbohydrate metabolism. In: Fructose-2,6-bisphosphate (ed. Pilkis, S. J.), pp. 65–85. Boca Raton: CRC Press. van Schaftingen, E. and Hers, G. G. (1981). Inhibition of fructose 1,6 bisphosphatase by fructose 2,6 bisphosphate. Proc. Nat. Acad. Sci. 78, 2861–2863. Vroemen, S. F., van Marrewijk, W. J. A. and van der Horst, D. J. (1995). Stimulation of glycogenolysis by three locust adipokinetic hormones involves Gs and cAMP. Molec. Cell. Endocrinol. 107, 165–171. Waldbauer, G. P. and Friedman, S. (1991). Self-selection of optimal diets by insects. Ann. Rev. Entomol. 36, 43–63. Waldbauer, G. P., Cohen, R. W. and Friedman, S. (1984). Self-selection of an optimal nutrient mix from defined diets by larvae of the corn earworm, Heliothis zea (Boddie). Physiol. Zool. 57, 590–597. Walker, P. R. and Bailey, E. (1969). A comparison of the properties of the phosphofructokinases of the fat body and flight muscle of the adult male desert locust. Biochem. J. 111, 365–369. Wang, C. H., Gregg, C. T., Forbusch, I. A., Christensen, B. E. and Cheldelin, V. H. (1956). Carbohydrate metabolism in bakers’ yeast. I. Time course study of glucose utilization. J. Amer. Chem. Soc. 78, 1869–1872. Wang, G. M. and Haymet, A. D. J. (1998). Trehalose and other sugar solutions at low temperature: modulated differential scanning calorimetry (MDSC). J. Phys. Chem. 102B, 5341–5347.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
283
Wang, Z., Hayakawa, Y. and Downer, R. G. H. (1990). Factors influencing cyclic AMP and diacylglycerol levels in fat body of Locusta migratoria. Insect Biochem. 20, 325–330. Wannet, W. J. B., Hermans, J. H. M., van der Drift, C. and Op den Camp, H. J. M. (2000). HPLC detection of soluble carbohydrates involved in mannitol and trehalose metabolism in the edible mushroom Agaricus bisporus. J. Agric. Food Chem. 48, 287–291. Wasserthal, L. T. (1997). Interaction of circulation and tracheal ventilation in holometabolous insects. Adv. Insect Physiol. 26, 298–352. Weis-Fogh, T. (1964). Diffusion in insect wing muscle, the most active tissue known. J. Exp. Biol. 41, 229–256. Whim, M. D. and Evans, P. D. (1988). Octopaminergic modulation of flight muscle in the locust. J. Exp. Biol. 124, 247–266. Wiemken, A. (1990). Trehalose in yeast, stress protectant rather than reserve carbohydrate. Antonie van Leeuwenhoek 58, 209–217. Wiens, A. W. and Gilbert, L. I. (1965). Regulation of fat body metabolism by the corpus cardiacum in vitro. Science 150, 614–616. Wiggers, H. A. L. (1832). Untersuchung uber das mutterkorn, Secale cornutum. Justus Liebigs Annln. Chem. 1, 129–182. Wigglesworth, V. B. (1942). The storage of protein, fat, glycogen and uric acid in the fat body and other tissues of mosquito larvae. J. Exp. Biol. 19, 56–77. Wimer, L. T. (1969). A comparison of the carbohydrate composition of the hemolymph and fat body of Phormia regina during larval development. Comp. Biochem. Physiol. 29, 1055–1062. Winderickx, H., de Winde, J. H., Crauwels, M., Hino, A., Hohmann, S., van Dijck, P. and Thevelein, J. M. (1996). Regulation of genes encoding subunits of the trehalose synthase complex in Saccaromyces cerevisiae: novel variations of STRE-mediated transcription control? Molec. Gen. Genet. 252, 470–482. Wolschek, M. F. and Kubicek, C. P. (1997). The filamentous fungus Aspergillus niger contains two ‘differentially regulated’ trehalose-6-phosphate synthase-encoding genes, tpsA and tpsB. J. Biol. Chem. 272, 2729–2735. Womersley, C. Z. (1987). A re-evaluation of strategies employed by nematode anhydrobiotes in relation to their natural environment. In: Vistas on Nematology (eds. Veech, J., and Dickson, D. W.), pp. 165–173. Hyattsville: Society of Nematologists. Womersley, C. Z. and Higa, L. M. (1998). Trehalose: its role in the anhydrobiotic survival of Ditylenchus myceliophagus. Nematologica 44, 269–292. Womersley, C. Z., Wharton, D. A. and Higa, L. M. (1998). Survival biology. In: Freeliving and Plant-parasitic Nematodes (eds Perry, R. N. and Wright, D. J.), pp. 271– 302. Wood, F. E. and Nordin, J. H. (1980). Activation of the hexose monophosphate shunt during cold-induced glycerol accumulation by Protophormia terranovae. Insect Biochem. 10, 87–93. Wood, T. (1985). The Pentose Phosphate Pathway. pp. 3–16. New York: Academic Press. Woodring, J. P. (1984). The effects of starving versus fasting on blood composition in larval house crickets. J. Insect Physiol. 30, 251–255. Woodring, J. P., McBride, L. A. and Fields, P. (1989). The role of octopamine in handling and exercise-induced hyperglycaemia and hyperlipaemia in Acheta domesticus. J. Insect Physiol. 35, 613–617. Wyatt, G. R. (1967). The biochemistry of sugars and polysaccharides in insects. Adv. Insect Physiol. 4, 287–360.
284
S. N. THOMPSON
Wyatt, G. R. and Kalf, G. F. (1956). Trehalose in insects. Fed. Proc. 15, 388. Wyatt, G. R. and Pan, M. L. (1978). Insect plasma proteins. Annu. Rev. Biochem. 47, 779–817. Xie, G. and Timasheff, S. N. (1997). The thermodynamic mechanism of protein stabilization by trehalose. Biophys. Chem. 64, 25–43. Yaginuma, T. and Yamashita, O. (1979). NAD-dependent sorbitol dehydrogenase activity in relation to the termination of diapause in eggs of Bombyx mori. Insect Biochem. 9, 547–553. Yaginuma, T. and Yamashita, O. (1980). The origin of free glycerol accumulated in diapause eggs of Bombyx mori. Physiol. Entomol. 5, 93–97. Yamashita, O. (1983). Egg diapause. In: Endocrinology of Insects (eds. Downer, R. G. H., and Laufer, H.), pp. 337–342. New York: A.R. Liss. Yamashita, O. and Suzuki, K. (1991). Roles of morphogenetic hormones in embryonic diapause. In: Morphogenic Hormones in Arthropods (ed. Gupta, A. P.), pp. 82–128. New Brunswick: Rutgers University Press. Yamashita, O., Hasegawa, K. and Seki, M. (1972). Effect of the diapause hormone on trehalase activity in pupal ovaries of the silkworm Bombyx mori L. Gen. Comp. Endocrinol. 18, 515–523. Yamashita, O., Yaginuma, T. and Hasegawa, K. (1981). Hormonal and metabolic control of egg diapause of the silkworm, Bombyx mori (Lepidoptera: Bombycidae). Entomol. Gen. 7, 195–211. Yanagawa, H. (1978). Tissue distribution, purification, and properties of multiple forms of hexokinase in the silkworm, Bombyx mori. Insect Biochem. 8, 155–158. Yoshinaga, K., Yoshioka, H., Kurosaki, H., Hirasawa, M., Uritani, M. and Hasegawa, K. (1997). Protection by trehalose of DNA from radiation damage. Biosci. Biotech. Biochem. 61, 160–161. Zachariassen, K. E. (1985). Physiology of cold tolerance in insects. Physiol. Rev. 65, 799–832. Zachariassen, K. E. (1991). The water relations of overwintering insects. In: Insects at Low Temperature (eds. Lee, R. E., and Denlinger, D. L.), pp. 47–63. New York: Chapman and Hall. Zachariassen, K. E. and Lundheim, R. (1992). The endocrine control of insect cold hardiness. Zool. Jb. Physiol. 96, 183–196. Zanotto, F. P., Raubenheimer, D. and Simpson, S. J. (1996). Haemolymph amino acid and sugar levels in locusts fed nutritionally unbalanced diets. J. Comp. Physiol. 166B, 223–229. Ziegler, R. (1984). Metabolic energy expenditure and its hormonal regulation. In: Environmental Physiology and Biochemistry of Insects (ed. Hoffman, K. H.), pp. 95–118. Heidelberg: Springer-Verlag. Ziegler, R. (1991). Changes in lipid and carbohydrate metabolism during starvation in adult Manduca sexta. J. Comp. Physiol. 161B, 125–131. Ziegler, R. and Schulz, M. (1986). Regulation of carbohydrate metabolism during flight in Manduca sexta. J. Insect Physiol. 32, 997–1001. Ziegler, R., Eckart, K. and Law, J. H. (1990). Adipokinetic hormone controls lipid metabolism in adults and carbohydrate metabolism in larvae of Manduca sexta. Peptides 11, 1037–1040. Ziegler, R., Eckart, K., Jasensky, D. and Law, J. H. (1991). Structure-activity studies on adipokinetic hormones in Manduca sexta. Arch. Insect Biochem. Physiol. 18, 229–237. Zitnan, D., Kingan, T. G. and Beckage, N. E. (1995). Parasitism-induced accumulation of FMRFamide-like peptides in the gut innervation and endocrine cells of Manduca sexta. Insect Biochem. Molec. Biol. 25, 669–678.
TREHALOSE – THE INSECT ‘BLOOD’ SUGAR
285
Zitnan, D., Sauman, I. and Sehnal, F. (1993). Peptidergic innervation and endocrine cells of insect midgut. Arch. Insect Biochem. Physiol. 22, 113–132. Zlatkis, A. and Kaiser, R. E. (1977). HPTLC High performance thin-layer chromatography. New York: Elsevier. Zudaire, E., Simpson, S. J. and Montuenga, L. M. (1998). Effects of food content, insect age and stage in the feeding cycle on the FMRFamide immunoreactivity of diffuse endocrine cells in the locust gut. J. Exp. Biol. 201, 2971–2979.
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Index Acyrthosiphon pisum 76, 89–90, 98, 100, 102, 104, 107, 110, 113–16, 118–19, 121 Aedes aegypti 18, 34, 47, 229, 257 Aedes taeniorhynchus 231 aerobic respiration 241 AeSCP-2 47 algae 4–6 Alnus glutinosa 9 Alnus pubescens 87 amino acids catabolism 116 contribution of essential and non-essential to honeydew 102 functions of 116–17 gluconeogenesis from 228–30 in phloem sap 86, 102–4 uptake by aphids 103 ammonia as nitrogen excretory product of aphids 117 animal sterols 9 Antheraea polyhemus 207 Anthonomus grandis 33 aphid morph 120–1 aphid saliva 91–2 aphid-induced changes to phloem sap 85–7 Aphidius ervi 121 aphids acquisition of nutrients from symbiotic micro-organisms 107–12 allocation of nutrients to embryos 117–18 amino acid uptake 103 ammonia as nitrogen excretory product 117 assimilation efficiency of nutrients 93–4 bacterial symbiosis 118–20 chemical transformations of ingested food in gut 94 ADVANCES IN INSECT PHYSIOLOGY VOL. 31 ISBN 0-12-024231-1
chemical transformations of sucrose in gut and osmoregulation 100–1 condition of stomach feeding from chenopods and other plants 106–7 contribution of essential amino acids and non-essential amino acids to honeydew 102 contribution of water flux to osmoregulation 101 diet 76–87 essential amino acids in 123–4 exceptional features of diet 74 fate of nutrients acquired by 112–21 feeding rates 89 food choice 122–3 food processing in alimentary tract 92–107 food processing studies 93–4 future directions 121–8 gut enzymes 99–100 ingested allelochemicals 104–7 integration of bacterial symbiosis into nutritional physiology 123–5 intimate symbiosis with micro-organisms 74 metabolic fate of acquired nutrients 112–17 microbiology 107–9 mouthparts and foregut 88 nitrogen nutrition 123 nutrient allocation patterns 118–20 determinants of 120–1 nutrient utilization regulation 122 nutritional physiology 73–140 parasitization effects 121 physiological fate of acquired nutrients 117–20 processing of ingested nitrogenous compounds 102–4 processing of ingested sugars 97–101 sterols in 117 Copyright # 2003 Elsevier Ltd All rights of reproduction in any form reserved
288
structural organization of alimentary tract 94–7 see also phloem sap Aphis fabae 106, 109–10, 116 Aphis gossypii 115, 127 Aphis jacobaea 105 Apis mellifera 20, 56, 215, 235 apoplastic loading 78–9 Apterygota 24–33 Arabidopsis 221 Arabidopsis thaliana 85 arachidonic acid 242 Baccharis viminea 8 basiconic sensilla 161, 170, 175 sensory neurones from 166 behavioural responses to chemosensory stimulation 180–2 of locusts 185 Bemisia argentifolii 104, 127 Betula pendula 87 BGL2 85 bimodal chemo/mechanosensory sensilla 160 Blaberus discoidalis 235, 241–2 Blaptica dubia 233–4, 237, 243 Blattella germanica 13 Boettcherisca peregrina 235 Bombyx mori 19, 22, 34, 44, 46, 52, 226, 231, 235–9, 241–2, 246–7 Brachycera 18 Brachymeria lasus 20 Brassica napus 80–1 Brevicoryne brassicae 106 Buchnera 74, 76, 107–8, 114, 117–21, 124–5 integration of production of essential amino acids into aphid nutritonal physiology 111–12 nutritional contribution to aphids 109–12 production of essential amino acids by 109–11 Calliphora erythrocephala 215, 245 Calliphora nigribarbis 235–6 Calluna vulgaris 9 Camponotus compressus 45 carbohydrates 112–15 in phloem sap 79–80 carcinogenesis 55 catabolism 236–41
INDEX
Cerataphini 128 chemoreceptors, mechanisms affecting 156 chemosensation, peripheral sensory physiology of 155 chemosensory afferents 149, 162, 175 chemosensory coding 147–58, 180–92 chemosensory mapping 168–70 chemosensory neurones 160, 191 response 155 sustained stimulation 156 chemosensory processing pathways 179 chemosensory response nutrient-specific modulation 157 variation in 156–8 chemosensory sensilla see contactchemosensory sensilla; sensilla chemosensory stimulation, behavioural responses to 180–2 chemosensory systems components of 144 irreversible changes 157 peripheral sensory physiology 143 Chenopodiaceae 106 Chenopodium album 106 Chilo supressalis 231 cholesterol 52, 117 conformational perspective 39, 41 in developing embryos 53 roles for 42 transfer to eggs 52 Chrysopa carnae 33 Coccinella septempuncta 33 Coleoptera 9 contact-chemoreception 143, 146 contact-chemoreceptor sensilla, sensory neurones from 165 contact-chemosensory coding, hypotheses 150–6 contact-chemosensory information 155 contact-chemosensory neurones basic functioning 151 responses of 149–50, 152 contact-chemosensory sensilla 164 sensory neurones from 162–4 sensory neurones innervating 167–8 see also sensilla contact-chemosensory system, dynamic role 158 Crambus tricestus 46
INDEX
Culex pipiens 18 cycloartenol 52 defences against insects 84–5 Delia antiqua 252 Delia brassicae 18 Delphacidae 127–8 Dermestes vulpinus 2 deterrence-coding neurones 171 deterrent (or ‘D’) neurone 150 diapause hormone 247–8 Diatraea grandiosella 44 dietary sources 4–9 dietary sterols 26–32, 41 Diptera 18 Diuraphis noxia 86 Drepanosiphum platanoidis 120 Drosophila 48, 50–1, 54, 164, 170 Drosophila maculatus 18, 45 Drosophila melanogaster 18, 34, 38, 50, 223, 246–7, 252 Drosophila mojavensis 34 Drosophila nigrospiracula 34 Drosophila pachea 18–19, 55 Dysdercus peruvianus 99 Dytiscus 45 EDTA exudation 79–81, 86, 123 electrical penetration graph (EPG) 89–90 Embden–Meyerhof glycolytic pathway 236 Endopterygota 13–20, 33–5 EPG atasets 122 Epiblema scudderiana 231, 233, 239 Epilachna varivestis 33, 37 essential amino acids in aphids 123–4 production from Buchnera 109–11 Euceraphis betulae 87 Eulachnus brevipilosis 96 Eurostra solidaginis 231, 233, 238–41 Euryocotis floridana 45, 49–51 Exopterygota 12–13, 24–33 exteroceptors, innervation 161 FMR famines 258 Fomes applanatus 7 fructose-1,6-bisphosphatase 230, 233 fructose-1,6-bisphosphate 234, 237–8 fructose-2,6-bisphosphatase 232, 244 fructose-2,6-bisphosphate 233–4, 237
289
fructose-6-phosphate 226, 229 fructose phosphate cycle 232–4 fungal sterols 6–7 GABA 119–20, 174, 186 galactose 115 gamma-amino butyric acid (GABA) 119–20, 174, 186 Gibberella fujikaroi 7 gluconeogenesis from amino acids 228–30 from glycerol 231 glucose concentrations in select insect species 217–19 trehalose biosynthesis from 220–3 glucose-6-phosphatase 229–30 glucose-6-phosphate 223, 227 glucosinolates 105–6 glyceraldehyde-3-phosphate 226 glycerol, gluconeogenesis from 231 Glycine max 81 glycogen degradation 241–4 trehalose biosynthesis from 231–2 glycogen phosphorylase 240 glycogen synthase 240 glycolysis 236–41 Glyphinaphis bambusae 115 Grammia genura 152 Gryllodes sigilatus 45 Gryllulus domesticus 13 gustatory information 153 gustatory processing 147–8 gustatory receptor proteins 151 gustatory signals, processing 170–80 H. virescens 51 haemolymph metabolites 214–20 haemolymph sugar levels 215 haemolymph transport 48–9 haemolymph trehalose concentration 215, 224 effects of physiological condition and experimental treatments 217–19 hedgehog (Hb) gene family 3 Helicoverpa zea 3, 19, 55–6, 253, 257–8 Hemiptera 20, 25, 126 Hieroglyphus nigrorepleteus 45 high-performance thin layer chromatography (HPTLC) 214
290
high-pressure liquid chromatography (HPLC) 214 honeydew ammonia in 116 contribution of essential and non-essential amino acids 102 honeydew sugars 97 hormonal regulation of metabolism 241–8 Hyalophora cecropia 221–2, 231, 235 hydrogen bonding in trehalose dihydrate 212 Hydrophyllum capitatum 8–9 5-hydroxytryptamine 116 Hymenoptera 9, 20, 34 Hypera postica 33 hypertrehalosemic hormone 241–4 Hypochaeris radicata 9 hypotrehalosemic activity 245–7 HzSNPV 56 inorganic substances in phloem sap 83–4 insecticides, insect resistance 57 intersegmental interneurones 178 intracellular transport 47–8 Juniperus uthaensis 8 koinobiont parasitoids 121 Larinus 206 leg motor neurones, responses of 185–8 Lepidoptera 9, 19 Leptinotarsa decemlineata 44 Leucophaea maderae 248 lichens 4–6 Lipaphis erysimi 106 lipids 113 metabolism, trehalose interactions with 248 in phloem sap 80–1 Lobus glomeratus 169 local circuit neurones 173 local circuits 170–80 in avoidance and acceptance behaviours 190 in metathoracic ganglion 187 motor output of 179–80 local interneurones 172 local leg avoidance reflex 181 Locusta migratoria 12, 24, 38, 53, 152, 171, 226, 243
INDEX
locusts behavioural responses 185 metathoracic ganglion 180–92 significance and general applicability of model 188–92 Lucilia sericata 2 Lupinus angustifolius 105 Macrosiphum euphorbiae 114 Mamestra brassicae 235, 257 Manduca sexta 2–3, 20, 34, 46–7, 49, 51, 55, 158, 167, 169, 184, 187–8, 209, 223–4, 227–32, 238, 242, 245, 252, 256, 258 Manulea replana 6 mechanoreception 146 mechanosensory afferents 162, 164 mechanosensory neurones 160 mechanosensory processing pathways 179 mechanosensory receptors 175 mechanosensory sensilla 162 metathoracic ganglion 162, 173 local circuits in 187 of locust 180–92 Microplitis demolitor 35 mitochondria 125 modality-specific segregation 164–7 Monema flavescens 239 motor output of local circuits 179–80 Musca autonmalis 216 Musca domestica 19, 34, 38, 50, 235 Myzus persicae 25, 85, 89, 95, 102, 104–6 Neisseria polysaccharea 99 Neodiprion pratti 34 neuronal signals 180 Nilaparvata lugens 104, 115 nitrogenous nutrients in phloem sap 81–3 NMR spectroscopy 227 nuclear magnetic resonance spectroscopy (NMR) 214, 227 octopamine 244 Oestroidea 19 olfactory coding 148–9 olfactory neurones 148 olfactory receptors 146 olfactory sensitivity 158 orthoperoid groups 158
INDEX
Otiorhynchus sulcatus 44 ovipositor valves 164 Pachycerpoideus vindemiae 20 Papilio machaon 239 parasitization effects in aphids 121 pentose phosphate pathway 224–7, 236 peripheral sensory physiology of chemosensation 155 Periplaneta americana 45, 221–2, 235–6, 243–4, 248 pH effects 216 phagostimulatory chemicals 156 phagostimulatory coding neurones 171 phagostimulatory power 153–4 phagostimulatory sensory neurones 153 Philosamia cynthia 239, 249 phloem sap 73–87 acquisition by aphids 87–92 amino acids in 86, 102–4 aphid-induced changes to 85–7 carbohydrates in 79–80 chemical protection 84 comparative physiology of feeders 125–8 composition 79, 123 diversity of phloem sap-feeding animals 125–7 impact of lesion-inducing aphids on 87 ingestion 87–90 inorganic substances in 83–4 insects feeding on 75 lipids in 80–1 loading 78 molecular physiology of symbiosis between phloem-feeding insects and micro-organisms 127–8 nitrogenous nutrients in 81–3 proteins in 104 reason for studying nutritional physiology of phloem sap-feeding insects 128–30 sampling 79 sterols in 80–1 transport 76–9 phloem sugars 74 Phormia 164, 170–1 Phormia regina 215, 221–2, 235, 245 6-phosphofructo-1-kinase 232–4, 237–9, 244 6-phosphofructo-2-kinase 232 phospholipids 113
291
Pieris brassicae 226, 231, 243 Pinus sylvestris 9 plant galls 87 plant sterols 7–9, 55, 81 plastids 125 Plutella xylostella 19, 54, 57 Polygonum sp. 8 polyols formation of 236–41 synthesis, hormonal regulation during cold-hardening 247 Populus fremontii 9 Populus nigra var. italica 9 protein stabilization during osmotic and thermal stress 251–3 protein synthesis 116 proteins in phloem sap 104 Protophormia terranovae 239 Pterocallis alni 87 Pyrausta nubilalis 19 pyrrolizidine alkaloids 105 Rhodnius prolixus 257 Rhopalosiphum padi 86, 91 Ricinus communis 105 S. exigua 46 Samia cynthia 231 Schistocerca 24 Schistocerca americana 12, 20, 24, 42–3, 45, 155–6 Schistocerca gregaria 45, 152, 207, 221, 237 Schizaphis graminum 25, 76, 86, 107, 111 Sciphithis obscurus 44 Senecio jacobea 105 senecionine-N-oxide 105 sensilla 144–5 locations and behavioural hierarchies 144–7 number of 157–8 sensory neurones 178 from basiconic sensilla 166 central projections 159–60 from contact-chemoreceptor sensilla 165 from contact-chemosensory sensilla 162–4, 167–8 from tactile hairs 160–3 sensory receptors 172 sensory signals, processing 172–8
292
Sericesthis geminata 44 sieve elements, stylet penetration to 90–1 sieve tube exudate proteins (STEPs) 82–3 Sitobion yakini 79 sitosterol 52 Solanum tuberosum 82 somatotopic mapping 167–8 sorbitol metabolism 241 spiking local interneurones 174, 176–7, 179, 186, 188 responses to different chemical solutions 182–4 Spodoptera frugiperda 235 Spodoptera littoralis 154, 254–5 Steinernema feltiae 56 sterol carrier protein-2 (SCP-2) 47 sterol metabolic capabilities 36 sterol metabolic constraints 56–8 sterol metabolic pathways 20–4 sterol metabolism 2–3, 9–38 patterns of 35–8 sterol nutrition 3 initial studies 2 and physiology 1–72 sterol reproductive physiology 52–4 sterol structure 2, 4–9 nomenclature 4 sterols absorption 44–7 allocation 54 in aphids 117 availability 2 dealkylation 23 dietary need 2 ecology 54–6 intake regulation 42–4 intracellular distribution 49–51 numbering systems 5 in phloem sap 80–1 physiology 3, 38–54, 58 profiles 26–32 survey of studies 10–11 taste 42–4 tissue distribution 47–54 transport 47–54 use 9–38 among insect orders 14–17 patterns of 35–8 stigmasterol diet 57 stylet penetration to sieve elements 90–1 Subsaltusaphis ornata 96
INDEX
sugars, analysis in haemolymph 213 Supella longipalpa 170 symplasmic loading 78 T. castaneum 46 taste model 189 neurobiology of 141 see also contact-chemoreception taste receptors 143 Tenebrio molitor 226, 229, 235, 243, 247 terminal palp sensilla 156 Thermobia domestica 24 total amount of sterol 39, 50 trehalase, hydrolysis 234–6 trehalose 114, 205–84 analysis 213–14 biosynthesis from dietary sugar 223–7 from glycogen 231–2 from gluconeogenesis 228–30 from glucose 220–3 chemistry 210–14 cryopreservation 249–51 discovery 206 enantiostasis 208–9 energy storage 248–9 functions of 209–10 homeostasis 208–9 hydrolysis 247–8 interactions with lipid metabolism 248 metabolism 208, 220–41 non-homeostatic regulation 209–10 physiological roles 248–59 prevention of radiation damage to DNA 251–2 principal role of 209 properties 210–13 regulation of feeding 253–9 role of 207 stress protection 249–53 structure 211 variable concentration 209 trehalose dihydrate, hydrogen bonding in 212 trehalose-6-phosphate synthase 222, 242 triacylglycerols 113 fatty acid composition 114 Trichiocamps populi 239
INDEX
Trichoplusia ni 232, 237 Trimerotropis saxatilis 6 Trogoderma granarium 18, 33, 37 Tuberaphis takenouchii 115 Tuberolachnus salignus 102 Varroa jacobsoni 56 Ventral Association Centre (VAC) 159, 162
293
Vicia faba 100, 102 vitrified membranes 251 Wigglesworthia sp. 126 Xenopsylla cheopi 20 Xiphydria maculata 34 yeast-like symbionts (YLS) 128
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