Structure, Function and Mode of Action of Select Arthropod Neuropeptides

Atta-ur-Rahman (Ed.) Studies in Natural Products Chemistry, Vol. Vol. 33 © 2006 Elsevier B.V. B.V. All rights rights reserved. ©

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STRUCTURE, FUNCTION AND MODE OF ACTION OF SELECT ARTHROPOD NEUROPEPTIDES GERD GADE and HEATHER G. MARCO Zoology Department, University of Cape Town, ZA-7701 Rondebosch, South Africa Dedicated to our parents who have always supported our scientific curiosity whole-heartedly and unconditionally. ABSTRACT: This overview summarizes important features of the majority of neuropeptide families that occur in two species-rich and widely-radiated arthropod taxa, the crustaceans and the insects. The neuropeptides may act as true neurohormones, which are released into the circulation, or as local neurotransmitter and/or neuromodulator. By comparing the primary structures of members of peptide families, the biosynthesis (including preprohormone structure and the peptidergic control of release), the structures of the receptors and transduction of the message via second messenger systems, the inactivation and the multiple functions of selected neuropeptides, we want to draw the reader's attention to the following main conclusions: 1) neuropeptides are important physiological regulators in arthropods; 2) neuropeptides can be structurally and functionally highly conserved in major arthropod groups (for example, proctolin, crustacean cardioactive peptide), or where peptide isoforms exist, there may be different scenarios: - (i) in the case of identical isoforms in insects and crustaceans, the peptide function may have changed in the two taxa (for example, red pigment-concentrating hormone affects pigmentation in crustaceans but mobilizes lipids in insects), - (ii) the isoforms are different in the two taxa and may have the same effect (for example, the ion-transporting peptide in insects and the crustacean hyperglycaemic hormone in crustaceans both play a role in osmoregulation), - (iii) the structurally different isoforms have different functions in the two taxa (for example, pigmentdispersing hormone affects pigmentation in crustaceans, whereas pigment-dispersing factor affects circadian rhythmicity in insects.

INTRODUCTION The phylum Arthropoda contains a number of subphyla, such as Chelicerata (scorpions, spiders and mites), Myriapoda (centipedes and millipedes), Crustacea (crabs, shrimps and woodlice) and Hexapoda (springtails, bristletails and insects).' The most conspicuous and wellknown members of the Arthropoda are, undoubtedly, the crustaceans and

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the insects. Even laymen recognize them and are fascinated by certain morphological features, behaviours and physiological events that are associated with these arthropods. For the specialist the debate is still ongoing as to whether crustaceans and insects are phylogenetically related in a sister group relationship; more and more evidence has recently accumulated from molecular data, ultrastructure of brain and eye, and from neurogenesis to support such a relationship [1-4]. Crustaceans and insects have diversified enormously, and scientific research has, thus, only been conducted on a small number of extant species. The crustaceans are divided into up to 6 classes [5] of which the Malacostraca is the largest and the one that contains the better known crustacean species that occur in the sea, freshwater and on land. Apart from the isopods, research on neuropeptides have mainly been conducted in decapod crustaceans, therefore, this review will be limited to the Decapoda, which contains the most familiar and widespread crustaceans such as shrimps, prawns, crabs, crayfish, spiny lobsters and lobsters and excludes the isopods and amphipods. The decapod crustaceans are a rich source of proteins and famous for their exceptional taste that has made them such a sought-after culinary delicacy. Decapod crustaceans, therefore, form the hub of commercially lucrative seafood industries the world-over. They are, however, also of ecological significance, chiefly dominating the marine and freshwater habitats with only a few terrestrial species. Insects, on the other hand, are the most successful animal group on land and, although insects form a substantial part of the diet of many human tribes, they are far more renowned for (a) their sheer abundance (at least 1 million species are described and a further estimated 3 - 5 million species have not been detected and/or described yet), (b) their striking beauty and (c) their beneficial aspects (e.g. pollination), as well as (d) their potential for disaster to mankind (e.g. crop damage, vector for the transmission of diseases). Insects and crustaceans are also well-known to the general public because of the spectacular changes that they undergo during their life-cycle, events such as metamorphosis and ecdysis; and who has never wondered about how these animals are able to adapt to environments with different salinities, or how they can change colour to camouflage themselves, or how they can use metabolites so efficiently during longdistance migrations or fast swimming? All these events, and more, are under the control of small, bioactive peptides. Because these regulatory peptides are synthesized in modified neurons, they are called neuropeptides.

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In this article we will review the existing literature on a selection of neuropeptides to find out whether the phylogenetically related decapod crustaceans and insects synthesize peptides of the same or very similar primary structures and if so, whether the neuropeptides are employed for the same function and whether this function is exerted by an identical pathway for their mode of action. Although such comparisons may shed some light on evolutionary aspects, by no means is this account meant to treat neuropeptides in a phylogenetic framework.

1. NEUROSECRETION Neuropeptides are peptidergic chemical messengers that are synthesized in specialized neurons and are released into the general circulation, which in insects and crustaceans is called haemolymph, to reach their target organ(s). Most neuropeptides are, in fact, hormones which control a number of physiological processes, hence, the neuroendocrine system represents a form of communication between cells, tissues and organs, other than the classical nervous and endocrine systems. Nervous control mechanisms act rapidly through synapses, releasing neurotransmitters into the synaptic cleft and generating action potentials of short duration; the classical endocrine (hormonal) control is slower acting but of longer duration since the hormones are released into circulation often a long distance away from the target organ and it takes some time before the hormones are degraded. Some neuropeptides do, however, also act as neurotransmitters and neuromodulators as evidenced by their distribution: the neuropeptide-synthesizing neurons may innervate a target organ directly, in addition to projecting to a neurohaemal organ, e.g. proctolinproducing neurons project into the pericardial organs (PO) of crustaceans, as well as to skeletal muscles, and proctolin has a positive modulating effect on neurons of the stomatogastric ganglion [6]. 1.1. Neuroendocrine complexes in insects Most of the endocrine processes in insects are controlled by neuropeptides. The neuroendocrine system in insects is comprised of neuropeptide-synthesizing neurons located in the cerebral ganglia (brain),

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specifically, in the pars intercerebralis and the median and lateral parts of the protocerebrum, as well as neurons in the retrocerebral corpora cardiaca (CC) and ventral nerve cord ganglia (see Fig. 1A). The neuropeptide producing cells are usually arranged symmetrically, and their axons end in so-called neurohaemal organs that serve as the storage and release site of the neuropeptide hormones. The chief neurohaemal organs of insects are (a) the paired retrocerebral CCs, that store neuropeptides from the brain, the sub-oesophageal ganglion and the CC itself and release them into the aorta and, (b) the ventral perisympathetic organs that store neuropeptides made by neurons in the thoracic and abdominal ganglia (see Fig. 1) [7]. A

B

Medial neurosecretory cells

Sinus gland

Lateral neurosecretory cells Brain -

Head

X organ

Corpus cardiacum Corpus allatum

ephalohorax

Gut

Perisympathetic organ

Abdomen

Abdomen

Figure 1. Schematic diagrams of the neuroendocrine system in (A) insects and (B) crustaceans showing the chief location of neurosecretory cells and their neurohaemal release sites. (A) is modified after [10]; fB) is modified after r 1121.

1.2. Neuroendocrine complexes in decapod crustaceans In crustaceans, too, many physiological processes are under neurohormonal control, and in the early years of investigation, these processes were pin-pointed by experimental biological studies that

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involved ablation or implantation of specific tissues/organs, injection of crude extracts of tissues/organs and observations at macroscopic and microscopic levels [8]. In this way it came to light that eyestalks of decapod crustaceans house an important endocrine centre that exerted control over glucose metabolism, moulting, reproduction and epithelial pigmentation. Histology with light microscopy revealed a neuroendocrine complex in the eyestalk that is made up of the X-organ (XO), a cluster of large neurons in the medulla terminalis which is the site of neuropeptide synthesis, and the sinus gland (SG), a neurohaemal organ which is located between the medulla externa and the medulla interna and is formed by the collective axonal endings of the XO neurons, and which serve as the site for storage and release of the neuropeptides into circulation (Fig. IB). Other sources of neuropeptides in decapod crustaceans have been identified [9]: (a) the pericardial organs (POs), neurohaemal organs that lie in the venous cavity surrounding the crustacean heart; the axons ending in the POs arise from thoracic (including the suboesophageal) ganglia and from intrinsic somata; (b) the post-commissural organs, neurohaemal organs with intrinsic neurons and axons arising from the commisure that passes posterior to the oesophagus, connecting the circumoesophageal connectives; not much is really known about this system, and (c) various components of the central nervous system (CNS), such as cerebral (brain), sub-oesophageal, thoracic and abdominal ganglia (Fig. IB). 2. METHODS USED IN NEUROPEPTIDE RESEARCH A suite of methods has been successfully applied over the past few decades in the research on arthropod neuropeptides: the relevant methods are mentioned only briefly here with some examples relating to crustacean neuropeptides; the reader is referred to [10] for a more detailed description of the general methods employed in the isolation and characterization of insect neuropeptides. 2.1. Biological assays Biological assays are employed to (a) determine the physiological relevance of a neuropeptide and (b) to monitor or identify the

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neuropeptide of interest during the procedure of isolation from an extract of biological material. The functional characterization and identification of neuropeptides includes studies at: (i) the gross in vivo level, e.g. which functions are affected by experimental intervention, such as ablation of crustacean eyestalks or re-implantation of sinus glands; (ii) the more-refined in vivo level, e.g. injections of assumed physiological doses of a chemically pure neuropeptide and monitoring specific parameters, such as measuring changes in glucose concentration after injection of crustacean hyperglycaemic hormone; (iii) the indirect in vitro approach, e.g. monitoring the effect that a neuropeptide has on a specific organ where this organ plays an integral role in another process, such as the Y-organ bioassay in which the effect of moult-inhibiting hormone on moulting is studied by measuring the output of ecdysteroids; (iv) the direct in vitro approach, e.g. monitoring the direct effect a neuropeptide may have on an explanted organ, such as cardioacceleratory peptide on heartbeat or the effect of myokinins on isolated hindgut. Other physiological studies with the isolated neuropeptides include dose-responses, time-course, mode of action, peptide titres and effects during stress and different stages of development or reproduction; crossactivity studies at intraspecies, as well as interspecies levels. Before the advent of molecular biological techniques, more often than not, a neuropeptide was first functionally identified by means of biological assays and subsequently sequenced than the other way around, i.e. very few peptides were sequenced without some knowledge of its function. 2.2. Elucidation of neuropeptide structure Once it was established that certain organs/tissues were producing a neuroendocrine factor and the actions of such factors were roughly defined by the experimental biological approach (biological assays), studies focused on the structural elucidation of the neuroendocrine substances. First, the chemical nature of the neurohormones was demonstrated as peptidergic (because they could be inactivated by proteolytic enzymes) [8] and then several other methods were employed to characterize the neuropeptides, e.g. size estimation by using gel chromatography, determining the iso-electric point and investigating whether the neuropeptide is heat-resistant or labile. It lasted several

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decades after its initial functional characterization before the first neuropeptide hormone from invertebrates, the red pigment-concentrating hormone (RPCH), was chemically isolated and its primary structure determined [11]. These early methods, however, required the use of gram amounts of starting material (e.g. whole eyestalks) plus chemically complicated procedures to isolate the active neuropeptide. Amino acid sequencing by Edman degradation in the 1970s was, of course, an important breakthrough in the field of peptide chemistry. Improved methods of microanalytic peptide chemistry, such as isolation of peptides by high pressure liquid chromatography (HPLC), peptide synthesis, reliable mass spectrometry for determining molecular weights and advanced functions of mass spectrometry for peptide sequencing, as well as the use of defined peptidases all contributed to the structural identification of neuropeptide hormones from different insects and crustaceans. 2.3. Localization and quantification of neuropeptides Information on the primary structures enabled comparative endocrinologists to arrange peptides into structurally homologous peptide families; one could also generate very specific antisera for localization of the neuropeptides in cells, tissues and organs with the aid of immunocytochemistry, and for peptide quantification (titre determinations) by means of immunoassays (radioimmunoassay; RIA and enzyme immunoassays; EIA). Polyclonal antisera, raised against conserved structural units of a known peptide, can also be employed to monitor and identify similar peptides during the isolation and purification procedures of the latter. For example, antisera raised against the crustacean hyperglycaemic hormone (cHH) of the edible crab, Cancer pagurus, and the American lobster, Homarus americanus, were successfully used in an enzyme-linked immunoassay (ELISA) to identify cHH molecules in an extract of sinus glands from the South African spiny lobster, Jasus lalandii [12].

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2.4. Molecular biological techniques Conventionally, invertebrate neuropeptides were isolated, chemically characterized and accompanied by some functional analyses. Since the movement towards molecular biological methods in the late 1980s, however, information of DNA encoding peptides with structural homology to known neuropeptide hormone families have "flooded" the scene, and in most of these cases, physiological functions of the encoded peptide could only be guessed at since the peptide itself was not chemically isolated. The chief advantage of using these methods is that relatively small amounts of starting material are required to (a) provide information on the preprohormone structures, (b) allow sensitive expression studies of the specific mRNA by in situ hybridization, Northern blot analyses or polymerase chain reaction (PCR), and (c) there is the potential for recombinant production of neuropeptides for use in physiological studies. The latter can be beneficial for the synthesis of larger and more complicated neuropeptide structures, such as the cHH family peptides which have proven a big challenge for traditional chemical synthesis [13, 14]. The use of improved molecular biology techniques and the world-wide impetus of such studies, also resulted in the sequencing of the complete genome of selected organisms, including the insects, Drosophila melanogaster and Anopheles gambiae. This served as a launch pad for what is now known as "database mining" where putative G proteincoupled receptors, for example, can be identified and then cloned. Although this has provided lots of structural information on receptors for insect neuropeptides, the ligands for these receptors are sometimes not known, hence the term "orphan receptor" has been coined. Prior to these technical advances, the primary structure of only a few receptors for invertebrate neuropeptides was known, and receptors were identified in fractionated cell membranes by specific binding of radiolabeled ligands. Receptor-ligand interaction was also investigated indirectly by performing functional bioassays with analogues of a particular ligand (e.g. the lipid-mobilizing assays with the migratory locust to gather information about the receptor for the insect adipokinetic hormone, AKH) [10]. Several assays are now commonly used to study putative receptors of neuropeptides and to "de-orphanize" the receptors: the receptor is expressed in mammalian cell lines or in amphibian oocytes and is then

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exposed to different peptide ligands; binding between the ligand and the receptor is then monitored in various ways, e.g. by bioluminescence or an electrophysiological signal. With the bioluminescence assay, cells are transfected to express (i) the G protein-coupled receptor of interest, (ii) a promiscuous G protein (so-called because it can activate most G proteincoupled receptors by the phospholipase C pathway) that effects an increase of intracellular Ca2+ when stimulated, and (iii) a Ca2+-sensitive photoprotein, apoaequorin. Aequorin is a bioluminescence complex that was originally isolated from a jellyfish; this complex is made up of three components, viz. (i) apoaequorin which has Ca2+-binding sites and changes conformation to an oxygenase when these sites are occupied, (ii) a luminophore cofactor called coelenterazine, and (iii) molecular oxygen. The molecular oxygen oxidizes coelenterazine and the emission of a blue light is one of the products formed [15]. Thus, in the bioassay when the agonist binds to the G protein-coupled receptor, the G protein is stimulated and results in an intracellular increase of Ca2+; the latter combines with apoaequorin and the protein undergoes a conformational change into an oxygenase, coelenterazine is oxidized and a blue light is emitted and measured [15]. The electrophysiologically based receptor bioassay relies on measuring a change in the membrane potential of cells when a ligand binds to the receptor of interest. One way to do this is by co-injecting the receptor cRNA with murine GIRK (G protein-gated inwardly rectifying potassium channel) cRNA into defolliculated oocytes from the frog Xenopus laevis. The electrophysiological events in the oocyte are recorded by whole cell voltage clamping in a high potassium bathing buffer [16]. When an agonist binds to the receptor, an inward potassium current is induced which is then recorded [16]. To date, the complete sequence information of a crustacean genome has not been published; consequently, there is a wide gap in the available data on receptors for crustacean neuropeptides. 3. INSECT AND CRUSTACEAN NEUROPEPTIDES It is by no means surprising that structurally homologous neuropeptides have been identified in crustaceans and insects - they share, after all, a common ancestry. The functions of some of these neuropeptides have not been conserved in these arthropod groups and we discuss here a selection of neuropeptide families common to both. For each peptide family, we

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will describe the characteristic structural features of the peptide members, when it was first isolated and elucidated in insects and crustaceans, functional activities, localization of the peptide and its synthesis, including available information on the gene and preprohormone, as well as known peptidergic factors regulating the release of the neuropeptide. Information on receptors and their interaction with specific neuropeptide ligands, as well as the peptide mode of action will also be discussed where possible. This review will not provide details, except in a few cases, of the various isolation techniques, nor of the many biological assays used in the characterization of neuropeptides and/or their receptors, since most of the commonly used methods have been elaborated on in previous reviews on neuropeptides [10] or are briefly described in the Introduction above (Section 2). In the various tables provided in the next sections, we will only give structures of those insect peptides that are not shown in our previous overview [10]; for crustaceans, however, we will give all published structures because they are not as readily available in earlier reviews.

3.1. Colour change versus metabolite mobilization: the AKH/RPCH and PDH/PDF families of neuropeptides Already as early as the 1920s to 1940s it was demonstrated that extracts from certain glands of crustaceans and insects caused "blanching" in shrimps [17-21], i.e. the shrimp changes from a dark to a light colour. In crustaceans, colour change is effected by the movement of pigment granules in the integumental cells (chromatophores): when the granules are concentrated in the cytoplasm, blanching results; when the granules are dispersed, a darkening effect is obtained. Later, it was discovered that pigment movement in another cell type of crustaceans was also under hormonal control: the compound eye of arthropods consists of a number of ommatidia which is the smallest functional unit in the process of visual perception. Whereas the movement of the screening pigments in the reticular (photoreceptor) cells is mainly achieved by direct action of light, pigment movement in non-reticular ommatidial cells (either reflecting pigment cells and/or distal pigment cells) is controlled by neurohormones [22,23].

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3.1.1. The AKH/RPCH family After a number of attempts of purification and separation of the peptides that affect pigments in chromatophores of various crustacean species [24], it took until 1968 [25] to achieve a complete purification of the material from the eyestalks of Pandalus borealis. The decisive step in the purification scheme was the elution behaviour of the compound in water/butanol mixtures on Sephadex LH-20 columns. The complete structure elucidation was made possible by: (a) quantitative ammo acid analysis and measurements of ultraviolet absorption (both methods together establishing the equimolar amount of 8 amino acids), (b) the use of the newly developed high-resolution mass spectrometer which determined the N-terminus as pGlu-Leu-Asn, (c) the successful proteolytic cleavage of the molecule into two fragments of 3 residues (the already known N-terminus) and 5 amino acid residues, and (d) the sequencing of the latter by Edman-dansyl sequencing to reveal the Cterminal sequence Phe-Ser-Pro-Gly-Trp amide [11, 26]. Thus, in 1972 the structure of the first invertebrate neuropeptide, now known as PanboRPCH because of its concentrating effect on red pigment granules in crustaceans, was completely elucidated (see Table 1). Parallel to these developments, scientists who were interested in carbohydrate and lipid metabolism in insects were looking for substances that have similar actions to those of the well-known metabolic hormones of vertebrates, glucagon and insulin. The first report on the existence of a glucagon-like factor in insects came from Steele [27]. Extracts of CC from the American cockroach, Periplaneta americana, elevated the concentration of trehalose, a disaccharide, which is the main blood (haemolymph) sugar of insects. A few years later a different effect of extracts from the CC was reported in the desert locust, Schistocerca gregaria [28], and the migratory locust, Locusta migratoria [29]. Here, the concentration of lipids in the haemolymph was increased upon injection of CC extract, thus, a hyperlipaemie or adipokinetic effect was measured. After a number of purification attempts, a decapeptide, which is now called locust adipokinetic hormone I (Locmi-AKH-I) was isolated from 3000 corpora cardiaca by size-exclusion chromatography on controlled-pore glass and thin layer chromatography on silica gel [30]. Structure elucidation was achieved by a combination of enzymatic

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cleavage and mass speetrometry resulting in the primary sequence of a decapeptide [30]. Table 1. Primary structures of members of the AKH/RPCH family of peptides. Peptide name, code name:

Species

Crustaceans red pigment-concentrating hormone Panbo-RPCH Pandalus borealis and various other crustaceans

Sequence

Reference

pELNFSPGW-NH2

[11.31]

Insects adipokinetic hormone Locmi-AKH-I

Locusta migratoria

pELNFTPNWGT-NH2

[30]

Loemi-AKH-H Locmi-AKH-ffl Panbo-RPCH Eiysi-AKH Pyrap-AKH Tenar-HrTH

L. migratoria L. migratoria Nezara viridula Erythemis slmpllcicollis Pyrrhocoris apterus Tenthredo arcuata

pEQLNFSAGW-NHj pELNFTPWW-NH2 pELNFSPGW-NHa pELNFTPSW-NH2 pELNFTPNW-NH2 pELNFSTGWGG-NH2

[232,233] [35] [32] [234] [235] [236]

A complete list of sequences up to 1997 is available for insect members of this family [10,37].

The primary structures of Locmi-AKH-I and Panbo-RPCH are strikingly similar (Table 1). Both peptides are cross-active in the reciprocal system and the structural similarity also explains why crude extracts of insect CCs (which contain one or the other form of "AKH") cause blanching in shrimps (see above). The structural similarity was also the basis for classifying these peptides as members of a peptide family, viz. the AKH/RPCH family of peptides. Thus, the peptide family includes structurally related peptides which have diverse functions in the two main arthropod taxa. New developments in the analysis of small peptides have greatly advanced this field. For example, improvements in the techniques of isolation and sequencing have occurred along with the introduction of modern mass spectrometric methods for accurate mass determination and sequencing tasks as first achieved with the fast atom bombardment (FAB) mode, and later with matrix-assisted laser desorption/ionization (MALDI) and electrospray ionization (ESI) mass speetrometry. These new techniques, combined with the large amount of peptidic material typically stored in the CC, made it relatively easy to determine the primary structure of almost 40 different natural analogues in about 100 insect species (see Table 1 for some examples). Far fewer decapod crustaceans have been investigated but, interestingly, the structure of RPCH is highlyconserved among crustaceans. In all species from which it has been

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sequenced or characterized by chromatographic behaviour and amino acid composition data, only RPCH can be found of the AKH/RPCH family peptides in crustaceans [see 31]. RPCH is, however, also synthesized in an insect species, the bug Nezara viridula (belonging to the large taxon of Hemiptera), where it mobilizes lipids [32]. In insects, representative peptides of the AKH/RPCH family have been found in most orders [33, 34]. In a number of taxa, gene duplication has taken place and two or even three AKH peptides are found in one species [10, 35-37]. In contrast to crustaceans (only RPCH present), insects show a high degree of variability in isoforms of AKH peptides. Common characteristics of the family are: a chain length of 8 to 10 amino acids; the N-terminus blocked by pyroglutamic acid (pGlu); the C-terminus blocked by a carboxyamide; amino acids at positions 8 and 9 (when present) are Tip and Gly; most of the peptides are uncharged, but there are a few that have an aspartic acid at position 7; there are at least two aromatic acids present, at position 4 mostly Phe (but sometimes Tyr) and at position 8 (Tip), and a few peptides have a third aromatic acid either at position 2 (Tyr or Phe) or at position 7 (Tip). In addition to the post-translational modifications at the terminals, the hypertrehalosaemic hormone (HrTH)-I of the stick insect, Carausius morosus (Carmo-HrTH-I) is glycosylated [38]. The site of glycosylation is not the usual Ser/Thr (O-glycosylation) or Asn (N-glycosylation) but Tip. As in human ribonuclease [39], the hexose in Carmo-HrTH-I is very likely C-glycosidically linked to the C-2 atom of the indole ring of Tip. In some insects, notably Lepidoptera (e.g. the butterfly Vanessa cardui) the AKH is not completely processed from the prohormone and, thus, occur in a C-terminally extended form (..-Gly-Gly-Lys) together with the fully processed peptide Manse-AKH [40]. Unfortunately, no rigorous tests have been conducted as to whether this compound is biologically active or whether the measured biological activity is the result of a breakdown product. Another, as yet unidentified, modification apparently occurs in a hypertrehalosaemic neuropeptide of cicadas: in a number of species two peaks are always found when using HPLC separation methodology but the material of both peak fractions have the same mass and amino acid sequence [10, 37]. When the synthetic peptide was synthesized according to the sequence information, its retention time on HPLC coincided only with one of the two peaks derived from natural material. The detailed pathways of the biosynthesis of the two adipokinetic hormones from S. gregaria, including the characterization of the

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preprohormones, have been elucidated by direct protein chemical methodologies, as well as molecular cloning [41, 42]. Similar studies have been executed for AKH family members in other insect species and also for RPCH of crustaceans. There is a distinct mRNA encoding for each AKH precursor (up to 3 in L migratoria) [43] which is translated into the discrete precursor, the prepro-AKHs. The organization of the precursor is basically always the same for all AKH and RPCH peptides: a signal peptide is followed by the respective AKH sequence, followed by the Gly residue for amidation of the AKH, the dibasic processing site, and C-termmally, a "tail peptide" or "precursor-related peptide". The latter is very long (more than 70 ammo acids) in the crustaceans compared to the 28 to 46 residues in insects. There are almost no structural similarities between signal peptides and "tail peptides" of insects and crustaceans. No biological function is known for any of the "tail peptides". Studies on the biosynthesis of the AKHs from the desert locust have revealed a unique strategy: after cleaving off the signal peptide, the two independently translated monomers of the pro-Locmi-AKH-I (as an example), consisting of the sequence for Locmi-AKH-I and the Cys-containing "tail peptide", respectively, are oxidized to a unique precursor dimer forming a disulfide bond. Thereafter, the precursor is processed to the following products: two monomeric molecules of Locmi-AKH-I extended by Gly-Lys-Arg and one dimeric molecule of the precursor-related peptide. The extended Locmi-AKH is subsequently cleaved by a carboxypeptidase H-like enzyme, which removes first the Arg and thereafter the Lys residue. A peptidylglycine-a-amidating monooxygenase then produces, from this Gly-extended form, the amidated Locmi-AKH. For RPCH and also other insect AKHs it is not known whether dimers are formed. The processes described above take place in the intrinsic neurosecretory cells of the CC of insects. The localization of AKH peptides in the neurosecretory cells of the CC has been shown numerous times by immunocytochemistry, and in situ hybridization experiments have demonstrated that the signals for the mRNA of all three AKH preprohormones of the migratory locust are co-localized in the neurosecretory cells of the CC [43]. RPCH immunoreaetive cells are found in neurons of the XO but also in other neurons, including those projecting into the neuropil of the stomatogastric ganglion of the crab, Cancer borealis and the spiny lobster, Panulirus interruptus or neurons of the abdominal ganglion of Pacifastacus leniusculm [see 6].

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Peptidergic releasing factors for AKHs of the migratory locust have been reported. In vitro studies identified the locust tachykinin [44] (see Section 3.4.6) and the crustacean cardioactive peptide [45, 46] which also occurs in insects (see Section 3.4.2) to promote the release of all three AKHs. The peptides FLRFamide from the desert locust and the peptide FMRFamide (see Section 3.4.4) were shown to be inhibitors of AKH release in the migratory locust in vitro [47]. There are no convincing reports on peptidergic releasing factors for RPCH in crustaceans and the reader is referred to [24] for information on the possible action of aminergic control factors. Release of AKH in locusts was also shown upon flight and quantitated by using a biological assay [48]. Only recently, sufficiently sensitive methods were published to determine reliable AKH titres by a RIA [49], and an ELISA [50]. In both immunoassays the major hurdle to overcome to obtain consistent measurements were the processing of the haemolymph samples. These are cumbersome and time-consuming and prevent the methods from being used on a routine basis. With the RIA, the release of Locmi-AKH-I and Schgr-AKH-II of the desert locust was shown to increase 15- and 6-fold upon 5 min of flight [49]. Since the maximal released levels are between 3 and 1 pmol for the two AKHs, it is clear that only a small fraction of the stored AKH material is released when locusts fly for 30 min. No titre determinations of RPCH have been executed, although there may be an RPCH analogue that could be useful for successfully developing a RIA method: when the Phe residue at position 4 in RPCH is replaced by Tyr this can be iodinated with 125I; a Tyr4 RPCH compound is even 4-fold more active than RPCH itself [51]. Once released into the haemolymph, the peptides are prone to proteolytic breakdown. Whereas no data are available on RPCH, there are quite a number of publications that deal with the fate of various AKHs. Only a few studies are reliable, those in which the AKH peptide concentration was used at physiological concentrations of about 1-3 pmol. In migratory locusts, for example, the three AKHs are not associated with carrier proteins during transport in the haemolymph, they have short half-lives which are not only different for the three peptides but also different for each peptide during a flight period [52]. One of the peptidases involved in the process of AKH breakdown is an enzyme with actions and physical and kinetic properties closely resembling those of the mammalian endopeptidase24.11 [10].

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Structure-activity studies are an indirect method to investigate a possible interaction of AKH peptides with their receptor proteins. For various members of the AKH family a vast body of literature has accumulated, employing bioassays such as lipid mobilization in locusts and the tobacco hornmoth Manduca sexta, carbohydrate mobilization in various cockroach species, the activation of phosphorylase in the larvae of M. sexta, or the inhibition of fatty acid synthesis in the fat body of locusts [10, 33, 37, 53]. Contrary to most other invertebrate neuropeptides, the insect members of the AKH family do not have a "core sequence" which is essential for potency, however, the N-terminal pGlu residue and the Cterminal amide are important, as well as the aromatics at position 4 and 8. To achieve full efficacy, all amino acids are apparently important. There is also no superagonist found and also no inhibitor. Two noteworthy sets of structure-function studies on RPCH have been performed. The chemical approach using synthetically modified analogues of RPCH and testing their effect on pigment movement in the shrimp, Palaemon (=Leander) adspersus revealed, surprisingly, that the blocked N-and C-termini are not essential for biological activity, in contrast to the situation in insects. Moreover, the Tip residue is very important for interaction with the receptor; replacement of the Phe residue at position 4 by Tyr results in an agonist with 4-fold higher activity [26]. A second approach made use of some of the then-existing natural analogues of RPCH, namely the insect AKH members, and measured their effect on erythrophores of the crayfish Cambarellus shufeldtii [54]. The native Panbo-RPCH was always the most potent compound, and the insect Schgr-AKH-II, which differs from Panbo-RPCH only in the Thr at position 6, had about 80 % of that potency; all other compounds were markedly less active. The first report on an AKH receptor protein in insects mentioned specific binding studies with tritiated Manse-AKH on membrane fractions purified from the fat body of adult M. sexta [55]. A competitive receptorbinding assay was developed and the analogues tested, revealing again that almost each amino acid of the AKH molecule is equally involved in interaction with the receptor [56]. In crustaceans to date, it has only been shown that membrane proteins from brain tissue, thoracic ganglia and the abdominal nerve cord are apparently able to bind RPCH [57], but no receptor has been isolated. This will, however, be feasible now, because the latest studies on insect AKH receptors have reported on molecular biological methods which may also be applicable to finding the RPCH

85

receptor. AKH receptors have been cloned from the fruit fly, D. melanogaster and the silkworm, Bombyx mori [58, 59]. These G proteincoupled receptors have 7 membrane-spanning domains and are structurally related to receptors of the vertebrate gonadotropin-releasing hormone. Following functional expression of the D. melanogaster AKH receptor in Chinese hamster ovary (CHO) cells [58] or in frog oocytes [59], the receptor responded to the AKH peptide of D. melanogaster with the lowest EC50 value of all AKH peptides tested. As a caveat, however, one has to stress that these tests are not testing for true receptor binding but are heterologous "functional" bioassays and often use a promiscuous G protein, which does not allow anyone to test whether the ligand really signals through Ca +. Studies concerning the cellular signalling pathways, thus the mode of action of AKH/RPCHs, have been numerous in insects but only few in crustaceans. If we consider the classical action of insect AKHs, namely to elicit an increase of carbohydrates, lipids or proline in the haemolymph, the following signalling pathways are known [37, 60-62]: to achieve the efflux of trehalose from the fat body cells, glycogen phosphorylase has to be activated and for this, AKH activates a Gq protein after binding to the receptor and this activation leads to the stimulation of phospholipase C (PLC) and the production of the second messengers inositol trisphosphate (IP3) and diacylglycerol; IP3 is responsible for Ca2+ mobilization from internal stores; external Ca2+ plays an important role as well. In the migratory locust, stimulation of adenylate cyclase (AC) and therefore the second messenger, cyclic AMP, is additionally involved in the activation of glycogen phosphorylase. On the other hand, those AKHs that are finally responsible to increase the concentrations of diacylglycerols or the amino acid proline in the haemolymph, signal via a Gs protein and a subsequently stimulated AC produces cyclic AMP. Again, extracellular and intracellular Ca2+ (from IP3 insensitive stores) is necessary to stimulate lipase. Pigment aggregation in the shrimp Macrobrachium potiuna is thought to proceed via a similar route involving PLC, IP3 and Ca2+ as described above, however, the evidence for this is weak and mostly indirect [63]. That insect AKHs are truly multifunctional and have pleiotropic tasks will be evident from the following list that summarizes the various effects observed besides the control of mobilization of metabolites [10, 37, 53]: 1. stimulation of lipid oxidation by flight muscles

86

2. increase of the lipid-carrying capacity of lipoprotein carriers in the haemolymph 3. inhibition of protein synthesis; maybe even a specific effect on vitellogenin and, thus, on reproductive processes 4. inhibition of synthesis of RNA 5. inhibition of synthesis of fatty acids 6. stimulation of heart beat and certain other muscles, and locomotory activity in general 7. aiding the immune response in the migratory locust The list of functions corroborated for RPCH is smaller [31]: 1. stimulation of the release of methyl farnesoate from mandibular organs (MOs) 2. modulation of the rhythms of certain parts of the crustacean stomatogastric and swimmeret system 3. modulation of the crustacean photoreceptor cells. RPCH is not known to have any true metabolic effect in crustaceans, however, RPCH can (of course) elicit most of the actions of the insect AKHs in the appropriate insect recipient. Likewise, insect AKHs are potent to concentrate pigments in the appropriate crustacean species but to date, no report has shown that AKH can effect a colour change in insects. 3.1.2. The PDH/PDF family Early studies had shown that there are substances in the eyestalks of decapod crustaceans that have an influence on retinal pigments (see above). In 1971 a peptide that caused the dispersion of distal retinal pigment was purified from eyestalk extracts of P. borealis mainly by chromatography on CM-Sephadex [64]; the biologically active fraction was sequenced by Edman-dansyl sequencing and the compound turned out to be an octadecapeptide with an amidated C-terminus but with a free N-terminus [65] (see Table 2). Due to its function, the peptide was originally called light adapting distal retinal pigment hormone (DRPH). It is now better known as pigment-dispersing hormone (thus, Panbo-ccPDH), because it also translocates the pigments in the chromatophores centrifugally [66]. About a decade later, a second PDH was chemically identified from the eyestalks of the fiddler crab, Uca pugilator, a so-

87

called P-PDH which differs from a-PDH in six positions (see Table 2) [67]. To date, PDHs from 14 decapod crustacean species are known; there are some modifications of a- and P-PDHs occurring. It was later discovered that extracts from heads of insects were able to elicit a dispersion of pigments in the epidermis of eyestalk-less fiddler crabs; this bioassay was employed to isolate the active principle from the grasshopper, Romalea microptera - a modified P-PDH - via a number of complex chromatographic steps including partition-, gel filtration- and ion-exchange chromatography [68]. Since then, the structures of pigmentdispersing factors (PDFs) have been completely elucidated from cricket, stick insect and cockroach; and orthologues have been identified in the genomic cDNA databases of D. melanogaster [69] and the malaria mosquito, A. gambiae [70]. There are many conserved structural features in the PDH/PDF family; namely the chain length (18 ammo acid residues), the N-terminal Asn and C-terminal amidated Ala residues, the residues Ser, He, Asn, Ser and Leu at positions 2, 5, 6, 7 and 9; moreover, the substitutions occurring at positions 4, 8, 10, 12, 15 and 17 can all be explained by point mutations [31]. Interestingly, like AKHs in insects, PDHs in crustaceans have undergone gene duplication and up to three forms may be found in one species, for example two a-PDHs and one P~ PDH, are found in Pandalus jordani [31] (Table 2). A phylogenetic tree for PDH/PDF peptides that is constructed with the sequence data, suggests that P-PDH may be an "ancient molecule from which the PDHs and PDFs evolved as a highly conserved family of neuropeptides" [31]. There is nothing known about the actual biosynthesis of these peptides. Molecular biological studies, however, have identified the preprohormone sequence from some PDHs and PDFs [31]. The general organization is strikingly different to that of the AKH/RPCH precursor: the signal peptide is followed immediately by a precursor-related peptide of unknown function and variable sequence and length, a dibasic or tribasic cleavage site and the PDH/PDF octadecapeptide with a C-terminal Gly for amidation and a monobasic or dibasic cleavage site at the end of the open reading frame (ORF). Localization of cells producing PDHs/PDFs has been achieved by immunoeytochemistry. Mapping of PDH-immunoreactivity in a few crustacean species revealed not only association with neurosecretory cells but also with intemeurons suggesting an additional role of PDH as neurotransmitter or in neuromodulation [71]. In several insects, including

88 88

D. melanogaster, PDF is localized in a few specialized neurons which have their cell bodies in the optic lobe and some processes in the accessory medulla of the optic lobe, an area which is known to house the neurons of master circadian pacemakers responsible for daily locomotory rhythms [72, 73]. It is from these and a number of other observations that PDFs are thought to be the communicator of these pacemaker cells to spread the message of the biological clock, thus acting as peptidergic neurotransmitters [74]. The PDFs do not affect pigment movement in insects. Table 2. Primary structures of members of the PDH/PDF family of peptides. Species Sequence Peptide name, code name Crustaceans a-pigment-dispersing hormone Panbo-PDH Pandalus borealis, Pandalusjordani Panjo-PDH-III P.jordani Macro-PDH Macrobrachium rosenbergii [5-pigment-dispersing hormone Ucapu-PDH Uca pugilator, Callinectes sapidus, Cancer magister, Carcinus maenas, Pacifastacus leniusculus Procl-PDH Procambarus clarkii, Orconectes immunis, Orconectes limosus Penaz-PDH Penaeus aztecus, Penaeus vannamei Panjo-PDH-I P.jordani Calsa-PDH-II C. sapidus Penva-PDH-III P. vannamei Penja-PDH-I Penja-PDH-II

Penaeus japonicus P. japonicus

Insects Pigment-dispersing factor Drome-PDF Drosophila melanogaster

Reference

NSGMINSILGIPRVMTEA-NH2 NSGMINSILGIPKVMADA-NH2 NSGMINSILGIPKVMAEA-NH2

[31,65] [31] [31]

NSELINSILGLPKVMNDA-NH2

[31]

NSELINSILGLPKVMNEA-NH2

[31]

NSELINSLLGIPKVMNDA-NH2 NSELINSLLGLPKVMTDA-NH2 NSELINSLLGISALMNEA-NH2 NSELINSLLGLPKVMNDA-NH2

[31] [31] [31] [31]

NSELINSLLGIPKVMTDA-NH2 NSELINSLLGLPKFMIDA-NH2

[31]

NSELINSLLSLPKNMNDA-NH2

[31]

[31]

Only one PDF structure is included; a complete list is available [10, 31].

A diurnal and circadian pattern of staining in terminals of specific PDF-expressing neurons in D. melanogaster was found that was consistent with a hypothesis of daily release of PDF [75]. Whether peptidergic compounds cause release of PDHs/PDFs is not known, but the involvement of aminergic compounds has been implicated [24]. A quantitative ELISA with an antiserum against the R. microptera PDF has been developed. It was used to determine that the optic ganglion of this grasshopper is the richest source of PDF, but titres of PDHs/PDFs in the haemolymph of any species have not been quantified [31]. It is well-

89

known that the two Met residues at positions 4 and 15 in certain PDHs are prone to oxidation and that the Met-oxidized PDH loses a great deal of its activity, but no systematic study has been published on the inactivation or the half-life of any member of this peptide family [31]. A G proteincoupled receptor is suggested for PDFs, but such a receptor has not yet been identified. There are, however, a few facts known about the interaction of the PDH ligand with its putative receptor from structureactivity studies [71]: a chain length of 13 amino acids (loss of the first 5 amino acids from the N-terminus) is the minimum structure to elicit pigment dispersion, albeit very weakly active. Progressively extending the molecule towards the N-terminus restores more and more activity. Replacing the Arg residue at position 13 with an arginine diphenylglyoxal derivative results in an agonist that is 14-fold more active than the parent molecule; it is suggested that this may be due to increased resistance to/protection against proteolytic cleavage, but this is not proven experimentally. Only a few studies have been conducted on the signalling of PDH in crustaceans. It is suggested that pigment dispersion is achieved by the ligand binding to a Gs protein-coupled receptor, resulting in the activation of AC and the increase of the intracellular concentration of cyclic AMP which in turn activates a cyclic AMP-dependent protein kinase (PKA) [63]. This PKA has been shown to phosphorylate a protein; such a signalling mechanism is also known to elicit pigment dispersion in vertebrate cells [63]. The main functions of PDHs in crustaceans are [see 31]: 1. inducing pigment-dispersion in all kinds of chromatophores, but often with a different potency for the various types of chromatophores, and, if more than one form of PDH is present in one particular species, they may have different potencies on the same type of chromatophores. 2. eliciting light adaptive movement of screening pigment in distal eye pigment cells, perhaps also in the reflecting pigments of some species. A vast body of information, including molecular and genetic studies, has accumulated on the role of PDFs mainly in D. melanogaster, the house fly, Musca domestica and the Madeira cockroach, Leucophaea maderae. All these activities have to do with rhythmic behaviours and biological clocks and it is suggested that PDFs act directly on brain or optic lobe neurons as both input and output factors for the circadian clock

90

controlling daily locomotion [31, 72, 74, 76]. However, there are also reports that PDF immunoreactive neurosecretory cells are found in locust abdominal ganglia which have neurohaemal release sites and, thus, the synthesized material could act as a neurohormone in the haemolymph [73]. 3.2. The cHH peptide family For many years, scientists were aware that peptide factors from the eyestalks of decapod crustaceans regulated glucose metabolism, moulting and female reproduction. It was only much later when these peptides were isolated from the XO-SG complex and structurally elucidated, that it became apparent that the peptides controlling these diverse processes were structurally homologous to each other and, hence, warranted inclusion as members of one peptide family. 3.2.1. Crustacean hyperglycaemic hormone The first member of the cHH peptide family to be functionally and structurally characterized from crustaceans was the crustacean hyperglycaemic hormone (cHH) of the shore crab, Carcinus maenas [77]; cHH got its name from the first function ascribed to this neuropeptide, viz. when injected into the haemolymph of a crustacean, there is a consequential and significant rise in the concentration of glucose in the haemolymph that peaks around 2 hours after injection. This is a relatively straight-forward biological assay to perform; cHH does, however, also have other functions which will be outlined below. The crab cHH has 72 amino acid residues, is blocked at the N-terminus by a pGlu residue and by an amide at the C-terminus, and contains six cysteines that form three intramolecular disulflde bridges [77]. The amino acid sequence of other peptides with cHH activity was later published from a number of other decapod crustacean species (see Table 3). In the American lobster, (Infraorder: Astacidae), 4 cHHs were isolated which arise from 2 different genes and subsequent post-translational modification. This entails isomerization of the Phe residue to yield peptides with L- or D-Phe3 residues. The D-Phe3-cHH displays a higher potency and an extended hyperglycaemic effect [78]; this isomerization of

91

cHH also occurs in other other astacideans, such as crayfish. In crabs (Infraorder: Brachyura), two cHHs are always isolated from the sinus glands in different concentrations; they are the products of one gene and subsequent post-translational modification, viz. the formation of a pGlu at the N-terminus; the cHH which is present in minor quantities, however, has the non-cyclized glutamine (Gin1) residue yet both cHH isoforms have the same potency in a hyperglycaemic bioassay [79]. The dynamics of the N-terminally blocked cHH peptide versus the non-blocked cHH of C. maenas was investigated, both peptides were cleared from the circulation at equivalent rates and no distinct differences in degradation and activity could be found between the two [80]. Thus, it seems as if the cHHs are not less stable or more vulnerable to exopeptidase degradation by having a free N-terminus. Such an unblocked cHH is, indeed, the norm in some other decapod crustaceans (see Table 3), e.g. prawns (Infraorder: Penaeidae) and spiny lobster (Infraorder: Palinura) where the multiple forms of cHHs are all unblocked at the N-terminus although the Cterminus is amidated; these cHHs are products of different genes. Table 3 shows that there are distinct similarities in peptide structure between species of the same infraorder. This may possibly explain the observed results of early heterologous hyperglycaemic bioassays in which cHH elicited a hyperglycaemic response in a phylogenetic group-specific manner [81]. Functional activity of cHH (and perhaps also its observed group-specificity) may be conferred by the C-terminal sequence of the cHH peptide; indirect evidence for this with respect to biological function came from conspecific bioassays with two C-terminally truncated cHHs of Jasus lalandii: such truncated cHH peptides did not elicit hyperglycaemia [82]. Support for this hypothesis was recently provided with conspecific bioassays in which recombinant cHH from the Kuruma prawn Penaeus japonicus was used: the non-amidated cHH peptide showed low hyperglycaemic activity compared to that induced by the amidated recombinant cHH [83]. The secondary structure of the amidated recombinant cHH (as determined from circular dichroism spectra) also differed to that of the non-amidated cHH, indicating that the C-terminal amide may be significant in the folding of the molecule [83]. The elevation of glucose in the haemolymph after injection of cHH into crustaceans results from the hormone acting on its target tissues to mobilize glycogen stores. Significant binding of radiolabelled C. maenas cHH to crude membrane preparations from hepatopancreas (midgut gland), heart muscle and epidermis have been observed [84]. Receptors

92

for cHH and their binding affinity for homologous and heterologous cHHs were also studied using classical binding assays with plasma membranes purified from the hepatopancreas of C. maenas and the crayfish Orconectes limosus; species specificity was demonstrated in this study, suggesting that the previously observed group specificity of cHH with regards to biological activity, reflected co-evolution of both the hormone and its receptor [85]. One should caution, however, that only one species from the two infraorders were tested in the heterologous displacement. Eyestalk ablation (i.e. removal of cHH) results in hypoglycaemia (low concentration of glucose in the haemolymph). This hypoglycaemia is accompanied by an increase in glycogen content in different tissues (e.g. hepatopancreas, muscle, epidermis), an inactivation of phosphorylase and an activation of glycogen synthetase [86]. Injection of purified cHH into O. limosus caused an increase in cyclic nucleotide levels and inactivated glycogen synthetase in hepatopancreas [87] and in muscle [88]. Although both cyclic nucleotides are also involved in the mode of action of cHH, cyclic GMP and not cyclic AMP appears to be the relevant second messenger of cHH in O. limosus [86]. It has been reported that synthesis of cHH is not restricted to the XO in the eyestalks of crustaceans, but is also shown by immunocytochemistry to be localized to the suboesophageal ganglion and thoracic second roots in H. americanus [89], in the POs of C. maenas [90] and is transiently expressed in gut paraneurons of C. maenas where it is involved with water uptake to facilitate ecdysis [91]. Interestingly, the PO-cHH of C. maenas has a free C-terminus, its first 40 amino acid residues are identical to the SG-cHH and, not surprisingly, it displays no functional activity in the cHH bioassay [90].

Table 3, Primary structures of members belonging to the crustacean hyperglycaemic hormone family. Peptide name, code name

Species

Crustaceans Crustacean hyperglycaemic hormone Homam cHH-A1 Homarus americams (=HomamMrH) 1 Homam cHH-B H. americanus Probo cHH1 OrclicHH

Procambarus bouvieri, Procambarus clarkii, Orconectes limasus

1

CarmacHH-II2

Carcinus maenas

Canpa cHH-tf

Cancer paguna

BytthcHH*

Bythograea thermidron

Penja cHH-I

Penaeusjaponicus

PenjacHH-II

P.japonicus

Penja cHH-ffl

P.japonicus

Penja cHH-V

P.japonicus

Penja cHH-VI

P.japonicus

3

Penmo cHH-I

Penaeus monodon

Penmo cHH-tf

Sequence

Reference

pEVFDOACKGVYDRNLFKKLDRVCEDCYNLYRKPFVATTCREHCYSNWVFRQCLDDLL LSDVIDEYVSNVQMV-NH2 pEVFDQACKGVYDRNLFKKLNRVCEDCYNLYRKPFVIVTCRENCYSNRVFRQCLDDLL MIDVIDEYVSNVQMV-NH2 pEVFDQACKGIYDRRIFKKLDRVCEDCYNLYRKPYVATTCRQNCYANSVFRQCLDDLL LIDWDEYISGVQTV-NH2 pEVFDQACKGI YDR&I FKKLDRVCEDCYNLYRKPY¥aTTCRQNCYJ«JSVFRQCLDDLL LIDVLDEYISGVQTV-NH2 pEIYDTSCKGVYDR&LFNDLEHVCDDCYNLYRTSYVASACRSNCYSNLVFRQCMDDLL MMDEFDQYARKVQMV-NH2 pEIYDTSCKGVYDRGLFSDLEHVCDDCYNLYRNSYVASACRSNCYSNWFRQCMEELL MMDEFDKYARAVQMV-NH2 pEIYDRSCKGLYDRRLFSDLDHVCDDCYNLYRNSRVANACRENCYSNLVFRQCMEDLL LMDQFDKYARAVQTV-NH2 SLFDPSCTGVFDRQLLRRLGRVCDDCFNVFREPNVATECRSNCYNNPVFRQCMAYW PAHLHNEHREAVQMV-NHs SLFDPSCTGVFDRQLLRKLGRVCDDCFNVFREPNVAMECRSNCYNNPVFRQCMEYLL PAHLHDEYRLAVQMV-NH2 SLFDPACTGIYDRQLLRKLGRLCDDCYNVFREPKVATGCRSNCYHNLIFLDCLEYLI PSHLQEEHMAAMQTV-NHz LVFDPSCAGVYDRVLLGKLNRLCDDCYDVFREPDVATECRSNCFYNLAFVQGLEYLM PPSLHEEYQANVQMV-NH2 LVFDPSCAGVYDRVLLGKLNRLCDDCYNVFREPNVATECRSNCFYNLAFVQGLEYLL PPSLHEEYQANVQM¥-NH2 SLFDPSCTGVFDRQLLRRLSRVCDDCFNVFREPNVATECRSNCYNNEVFRQCMEYLL

[102,237] [237] [238,239] [240] [77] [79] [241] [242] [242] [242] [242] [242] [243]

ANFDPSCAGVYNRELLGRLSRLCDDCYNVFREPKVATECRSNCFYNPVFVQCLEYLI

[243]

P. monodon P. monodon

ANFDPSCAGVYNRELLGRLSRLCDDCYNVFREPKVATECRNNCFYNPVFVQCLEYLI PADLHEEYQAHVQTV-NHj SLFDPACTGIYDRQLLGKLGRLCDDCYNVFREPKVATGCRSNCYYNLIFLDCLEYLI PSHLQEEHMEALQTV-NH2

[243]

3

Penmo cHH-III

Penmo cHH-IV

[243]

93

P. monodon 3

P. monodon

Pensc cHH

Penaeus schmidtii

Meten cHH-A* Meten cHH-B*

Metapenaeus ensis

Macro cHH Jasla cHH-I

Macrobrachium rosenbergii Jasus lalandii

Jasla cHH-II

J. lalandii

ANFDPSCAGVYDRELLGGLSRLCDDCYNVFREPKVATECRSNCFYNSVFVQCLEYLI PADLHEEYQAHVQTV-NH2 ANFDPSCTGVYDRELLGRLSRLCDDCYNVFREPKVATECRSNCFYNPVFVQCLEYLI PADLHEEYQAHVGTV-NH2 SLFDPSCSGVFDRELLGRLNRVCDDCYNVFRDPKVAMECKSNCFLNPAFIQCLEYLL PEDLHEEYQSHVQVV-NH2 SLFDPSCTGVKDRELLGRLNRVCDDCYNVFREPKVATECRSHCFLNPAFIQCLEYII FEVLHEEYQANVQLV-NH2 AILDQSCKGIFDRELFKKLDRVCDDCYNLYRKPYVAIDCRRGCYQNLVFRQCIQDLQ LMDDLDEYANAVQTV-NH2 AVFDQSCKGVYDRSLFSKLDRVCDDCYNLYRKHYVATGCRRNCYGNLVFRQCLDDLM LVDVVDEYVASVQMV-NH2 AVFDQSCKGVYDRSLFKKLDVVCDDCYNLYRKPYVATGCRENCYSNLVFRQCLDDLM LVDVVDEYVSTVQMV-NH2

[243] [244] [245] [246] [247] [12] [82]

Moult-inhibiting hormone Procl MIH Probo MIH Carma MIH Canpa MIH Calsa MIH* Chafe MIH* Canma MIH* PenjaMIH Penmo MIH-I* Penmo MIH-II* Penva MIH*

Procambarus. clarkii Procambarus bouvieri, Carcinus maenas Cancer pagurus Callinectes sapidus Charybdisferiatus Cancer magister Penaeus japonicus Penaeus monodon P. monodon Penaeus vannamei

RYVFEECPGVMGNRAVHGKVTRVCEDCYNVFRDTDVLAGCRKGCFSSEMFKLCLLAM ERVEEFPDFKRWIGILNA-NH2 pEVFDQACKGIYDRAIFKKLELVCDDCYNLYRKPKVATTCRENCYANSVFRQCLDDLL LINVVDEYISGVQIV-NH2 RVINDECPNLIGNRDLYKKVEWICEDCSNIFRKTGMASLCRRNCFFNEDFLWCVHAT ERSEELRDLEEWVGILGAGRD RVINDDCPNLIGNRDLYKKVEWICEDCSNIFRNTGMATLCRKNCFFNEDFLWCVYAT ERTEEMSQLRQWVGILGAGRE RVINDDCPNLIGNRDLYKKVEWICDDCANIYRSTGMASLCRKDCFFNEDFLWCVRAT ERS S DLAQLKQWVTILGAGRI RVINDDCPNLMGNRDLYKKVEWICDDCANIYRITGMASLCRKDCFFNFDFLWCVRAT FRS FDMTQLKQWVRILGAGRI RVINDDCPNLIGNRDLYKRVEWICEDCSNIFRNTGMATLCRKNCFFNEDFLWCVYAT ERTEEMSQLRQWVGILGAGRE SFIDNTCRGVMGNRDYNKKVVRVCEDCTNIFRLPGLDGMCRNRCFYNEWFLICLKAN REDEIEKFRVWISILNAGQ SLTDGTCRGRMGNREIYKKVDRVCEDCANIFRLPGLEGLCRDRCFYNEWFLLCLKAA NREDEIENFRVWVSILNA SLTEGTCRGRMGNREIYKKVDRVCEDCANIFRLPGLEGLCRDRCFYNEWFLLCLKAA NREDEIENFRVWISILNA DTFDHSCKGIYDRELFRKLDRVCEDCYNVFREPKVATECKSNCFVNKRFNVCVADLR HDVSRFLKMANSALS

[101] [103] [99] [100] [248] [249] [250] [104] [251] [251] [252]

94

Penmo cHH-V3

Meten MIH-A*

Metepenaeus ensis

Meten MIH-B*

M. ensis

Macro MIH-A* Macro MIH-B*

Macrobrachium rosenbergii M. rosenbergii

Jasla MIH4

Jasus lalandii

Vitellogenesis-inhibiting hormone Homam VIH1 Homarus americanus Nepno VIH* 5

Probo VIH

Nephrops norvegicus Procambarus bouvieri

SYIENTCRGVMGNRDIYKKVVRVCEDCTNIFRLPGLDGMCRDRCFNNEWFLVCLKAA NRDDELDKFKVWISILNPGL FSIDYTCTGAMGNRDIYNKVSRVCDDCANIYRLPGLDGMCRNRCFNNFWFMICLRAA KREDEIDKFRVWISILNPGGAW RYLDDECPGVMGNRDLYEKVVRVCDDCSNIFRMNDMGTRCRKDCFYNVDFLWCVYAT ERHGDVDQLNRWMSILRAGRK RFLDDECRGVMGNRDLYEYVVRICDDCENLFRKSNVGSRCKKNCFYNEDFMWCVRAT ERTDELEHLNRAMSIIRVGRK RFTFD-CPGMMGQRYLYEQVEQVCDDCYNLYREEKIAVNCRENCFLNSWFTVCL QATMREHETPRFDIWRSILKA-NH2

[253] [106] [107] [107] [105]

ASAWFTNDECPGVMGNRDLYEKVAWVCNDCANIFRNNDVGVMCKKDCFHTMWFLWCV YATERHGEIDQFRKWVSILR ASAWFTNDECPGVMGNRDLYEKVAWVCNDCANIFRINDVGVKCKKDCFHNMDFLWCV YATERHGEIDQFRKWISILRAGRK pEVFDQACKGIYDRAIFKKLELV****YN******VATTCRENCYAN

[254]

RRINNDCQNFIGNRAMYEKVDWICKDCANIFRKDGLLNNCRSNCFYNTEFLWCIDAT ENTRNKEQLEQWAAILGAGWN RRINNDCQNFIGNRAMYEKVDWICKDCANIFRQDGLLNNCRSNCFYNTEFLWCIDAT ENTRNKEQLEQWAAILGAGWN QIFDPSCKGLYDRGLFSDLEHVCKDCYNLYRNPQVTSACRVNCYSNRVFRQCMEDLL LMEDFDKYARAIQTV-NH2

[256]

SFFDIQCKGVYDKSIFARLDRICEDCYNLFREPQLHSLCRSDCFKSPYFKGCLQALL LIDEEEKFNQMVEIL-NH2 SFFTLECKGVFDAAIFARLDRICDDCFNLFREPQLYTLCRAECFTTPYFKGCMESLY LYDEKEQIDQMIDFV-NH2 SNFFDLECKGIFNKTMFFRLDRICEDCYQLFRETSIHRLCKQECFGSPFFNACIEAL QLHEEMDKYNEWRDTL-NH2

[126, 128]

[116] [255]

Mandibular organ-inhibiting hormone Canpa MOIH-1

Cancer pagurus

CanpaMOIH-2

Cancer pagurus

Libem MOIH*

Libinia emarginata

[256] [122]

Insects Ion-transporting peptide Schgr ITP* Bommo ITP* Drome ITP*

Schistocerca gregaria, Locusta migratoria Bombyx mori Drosophila melanogaster

[130] [131]

95

Cys residues are indictaed in BOLD text. 'Present as L-Phe3 and D-Phe3 isoforms. 2Sequence of cHH-I differs only by Gin1 residue. 3Sequenced by mass spectrometry. 4 A gap has been introduced to align the Cys residues. 5Partially sequenced; unclear residues are shown as*. "Sequence deduced either in part or completely from the nucleotide sequence.

96

3.2.2. Moult-inhibiting hormone The second group of peptides that belong to the cHH peptide family are the moult-inhibiting hormones (MIH), so-called because it increases the interval between subsequent moults by exerting an inhibitory action on the Y-organ (YO), a cephalothoracic gland in which the moultpromoting steroid hormones (ecdysteroids) are synthesized [8]. The accepted, simplified paradigm for this is as follows: like all arthropods, crustaceans have a chitinous exoskeleton that must be shed and replaced by a larger exoskeleton in order for the animal to increase in size but, unlike insects, the moult cycle of crustaceans is under negative control and, only when the repressive hold of MIH on the YO is lifted, can the YO produce sufficient titre of ecdysteroids that leads to ecdysis. Confirmation of this paradigm has been provided in numerous cases where eyestalk ablations (thus, removal of the source of MIH) accelerated the moult cycle and enhanced the synthesis of ecdysteroids by the YOs [92] and by quantitative, moult-specific changes in MIH gene expression, as determined by Northern blots with eyestalk neural ganglia from the blue crab Callinectes sapidus at different stages of the moult cycle [93]. The latter result is in contrast to a similar study carried out with MIH mRNA from P. japonicus where no moult stage-specific fluctuation in MIH expression was observed [94]. Both studies have revealed, however, that MIH is expressed throughout the moult cycle. To date, however, the titre of MIH has not been determined in the haemolymph of any crustacean during all stages of the moult cycle, despite having MIHspecific antisera available; a chief reason for this is that the hormone is released in a pulsatile manner in minute quantities and has a short half-life of 5-10 min [95]. On the other hand, the titre of circulating ecdysteroids has been established in several decapod crustaceans (see for example [96]). On the basis of ecdysteroid measurements, a reliable in vitro bioassay was developed by Soumoff and O'Connor [97] to measure the reduction of ecdysteroid synthesis in YOs after exposure to crude extracts of SGs (or MIH). This assay was successfully used to characterize the first MIH to be sequenced [98]; the primary structure of this neuropeptide (see Table 3), isolated from SGs of C. maenas [99] was, surprisingly, similar to the neuropeptides involved in glucose metabolism, i.e. the cHHs. The structure of MIH isolated from another brachyuran crab, viz. the edible crab Cancer pagurus [100] showed that the sequences are very

97

highly related (around 80 % structural identity): both consist of 78 residues with a free N- and C-terminus and 6 cysteine residues in conserved positions that form 3 intrachain disulfide bridges (see Table 3). On the other hand, there is only around 30 % sequence identity between the brachyuran MIH and cHH peptides. In astacurans, a structurally distinctive MIH has, thusfar, only been reported in the crayfish Procambarus clarkii [101]; a peptide with MIH activity in H. americanus [102] and the Mexican crayfish Procambarus bouvieri [103] has also been reported on but, structurally, these neuropeptides are more characteristic of cHHs than MIHs (see Table 3). Peptides with MIH activity and which are structurally similar to the brachyuran MIHs have also been isolated and functionally characterized from sinus glands of P. japonicus [104] and J. lalandii [105]. Many more putative MIH sequences have been deduced from cDNA sequences of a variety of decapod crustaceans, notably brachyuran crabs and prawns (see Table 3). Although these deduced sequences show homology to MIH peptides, there is, in most cases, no additional evidence that they are indeed functional MIHs. One exception is the study in which the expression of a putative MIH of C. sapidus was investigated by Northern blots during the moult cycle [93]. In another study that tried to functionally characterize 2 putative MIH clones from the prawn Metapenaeus ensis, recombinant MIHs were produced in Escherichia coli and shown to increase the interval between subsequent moults when injected into M. ensis specimens [106]. There are other reports of possibly more than one MIH isoform being present in crustaceans: 2 MIH-like sequences have been deduced from cDNAs of the giant freshwater prawn Macrobrachium rosenbergii [107] and from the giant tiger prawn Penaeus monodon (see Table 3); it should be noted, however, that these putative MIHs have not yet been functionally characterized. There are several reports that cHH can inhibit the ecdysteroid synthesis by YO in vitro but that MIH cannot act as a cHH by increasing glucose concentration (see for example [108]). It was probably this multifunctionality of cHH that led to it being mistaken for MIH in certain instances. Why should cHH duplicate the role of another hormone during the moult cycle? The definitive answer to this question is still not known but specific receptors for cHH (along with MIH-specific receptors) have been demonstrated, by classical membrane binding studies, to be present on the YOs of intermoult brachyuran crabs [84]. Unlike cHH, radiolabelled MIH binds only to membrane preparations of YOs and not

98

to epidermal, heart muscle and hepatopancreas preparations [84]. Not surprisingly, MIH peptides from 2 other crab species could bind effectively to the C. maenas MIH receptor, which suggests that there is a high degree of conservation in the binding domains of the crab MIHs [84]. What is surprising, though, is that neither the density nor the affinity of the MIH and cHH receptors on the YO change significantly during the moult cycle of the crab [95], yet MIH and cHH have very little inhibitory effect in vitro on YOs at certain stages of the moult cycle (see for example [96]). This seems to suggest that the degree of YO inhibition may be independent of hormone titre and receptor expression, and further, that the control of signal transduction must lie downstream of the MIH receptor, thus the accepted paradigm of moult control is certainly oversimplified and possibly inaccurate [95]. The binding of MIH to its receptor on the YO cell membrane activates a signalling pathway that results in the inhibition of: (a) the cellular uptake of the ecdysteroid precursor (cholesterol), (b) protein synthesis and (c) the expression of steroidogenic enzymes [109]. Crustaceans are unique with respect to the negative control of ecdysteroid synthesis: after receptor-ligand interaction, there is an increase in cyclic nucleotide levels and this shuts down the YO cell's activity [110]. Although both cyclic AMP-dependent protein kinase (PKA) and cyclic GMP-dependent protein kinase (PKG) are present in YOs of O. limosus, only the activity of PKG is decreased by the removal of eyestalks, indicating that cyclic GMP is the second messenger of MIH [109]. Steroidogenesis is clearly enhanced by intracellular Ca2+ which arises from an influx from the extracellular space (via channels) and from the release of intracellular stores via IP3 [109]. Numerous investigations have shown that MIH functions independently of Ca2+ concentration, although its action can be eliminated by a high concentration of Ca2+ [110]. Few studies have looked at the expression of MIH apart from the classical XO-SG distribution, and the results are not all in agreement. Northern blots, PCRs and immunocytochemistry with antisera raised to recombinant MIH-B of M. ensis showed that MIH-B is expressed in the eyestalk (XO) and ventral nerve cord [106], whereas PCRs and Northern blots showed expression of a putative MIH only in the eyestalks of M. rosenbergii [107]. In C. pagurus, MIH-like transcripts were detected by Northern blotting only in XOs, whereas the more-sensitive nested PCR approach revealed that MIH could be amplified from optic nerve and the

99

ventral nerve cord [111]. Whether this non-XO production of MIH has any physiological role is still not clear. 3.2.3. Vitellogenesis-inhibiting hormone A third member of the cHH peptide family is also synthesized by neurons of the crustacean XO and has been named the vitellogenesisinhibiting hormone (VIH) or the gonad-inhibiting hormone (GIH) because of its negative influence on reproduction; this inhibitory peptide does not act in a species- or group-specific manner (for review see [112]. The identification of VIHs has been slow owing to a lack of reliable and fast bioassays [92]: in vivo assays in which ovarian growth or oocyte diameter is measured after injection of VIH, is only feasible with crustaceans that have short vitellogenic cycles, hence, in vitro assays which measure the endocytotic uptake of gold-labelled vitellogenin, or measures the endogenous synthesis of ovarian proteins by means of the incorporation of radiolabelled amino acids have been favoured. Heterologous bioassays with shrimps and prawns are most often used because these crustaceans have short and numerous reproductive cycles per year, the approximate vitellogenic stage of the ovary can be visually assessed through the opaque exoskeleton of the living animal and the reproductive/vitellogenic cycle is well documented (see for example [113]). Although the heterologous peptides have a significant effect at physiological doses, a homologous bioassay would provide unequivocal data/evidence as to whether the peptide is a true VIH in its own system. All VIHs sequenced to date from decapod crustaceans have been functionally characterized in heterologous assay systems: the first was isolated from H. americanus and tested in an in vitro assay with shrimps [114]. The VIH peptide consists of 78 amino acid residues, a free Nterminus but an amidated C-terminus and 6 Cys residues with 3 disulfide bridges (Table 3). It is around 53 % identical to MIH molecules and 25 % identical to cHHs [92]. The only other functionally characterized decapod VIH is a partially sequenced peptide that was isolated from P. bouvieri; this peptide inhibited the growth of prawn oocytes in vitro [115] but this activity was attained with an unphysiologically high peptide dosage and further, its structure resembles a cHH rather than the lobster VIH (Table 3). Recently, a cDNA sequence from the eyestalks of the Norway lobster Nephrops norvegicus was published [116]; the cDNA encodes a peptide

100

that is 96 % identical to the preproVIH of//, americanus [117]. The N. norvegicus VIH must, however, still be characterized to confirm its biological significance. To date, a characteristic VIH peptide has not been found in any brachyuran crab, spiny lobster, shrimp or prawn [see 113]. VIH activity, as gleaned from heterologous in vitro assays, is not associated with a unique peptide from the XO-SG complex of the spiny lobster J. lalandii [113] and the prawn P. japonicus [118], but was elicited by previously characterized cHHs, whereas MIH had only a neglible effect in both studies. In contrast, the lobster VIH displayed no hyperglycaemic activity and the lobster cHH did not have a VIH effect [119], although the lobster cHH clearly has MIH activity [102]. There is, unfortunately no report that a brachyuran MIH has been tested in a VIH assay. Interestingly, immunocytochemistry and in situ hybridization studies show that VIH is localized not only in female crustaceans but also in males and larvae [92]; the biological significance of this is still unknown but this distribution of VIH is a strong argument for changing the name of the peptide since its function may not be limited to the inhibition of vitellogenesis in female crustaceans.

3.2.4. Mandibular organ-inhibiting hormone The most recent member of the large cHH peptide family in crustaceans was functionally characterized as a mandibular organinhibiting hormone (MOIH) in brachyuran crabs [120]. The MO, which is located at the base of the tendon associated with the adductor muscle of the mandible, synthesizes methyl farnesoate (MF), a precursor of the juvenile hormone (JH) that occurs in insects, viz. JH III [121]. Earlier, observations were made that the removal of eyestalks leads to hypertrophy of the MO and more recently, it was shown that the circulating MF titres are increased after eyestalk ablation in three different crustacean species [121]. Thus, it was not surprising that the MOIH peptide was isolated from the SG of crustaceans. The biological assay used in identifying a MOIH peptide is based on the quantitative inhibition of MF synthesis by MOs in vitro, following incubations with the peptide. In this way, two MOIHs were identified in C. pagurus, isolated from SGs by HPLC and N-terminally sequenced [120]. These MOIHs are 78 residues long, have free N- and C-termini, three intrachain disulfide bridges and differ by only one amino acid substitution (Table 3).

101 101

Although there is 59 % sequence identity between C. paguras MIH and MOIH, the MIH is not active as an MOIH and the MOIH displays only limited activity in the MIH bioassay; furthermore, the cHH of this crab did not affect the MO in an in vitro MOIH assay [120], hi contrast, the 2 cHHs of C. maenas were equipotent at inhibiting the MOs in vitro (inhibition of around 83 %), whereas the MIH effected only 12.5 % inhibition [108]. hi another conspecific in vitro bioassay, the MIH and the 2 cHH peptides of O. limosus effected 80-89 % inhibition of MF synthesis [108], Peptides with MOIH activity have also been identified in the spider crab Libinia emarginata (Table 3) with an in vitro bioassay using dissociated MO cells; these peptides, however, also demonstrated cHH activity and are structurally more similar to the cHH than to the MOIHs of C. pagurus [122]. Criticism has been levelled against this study on the basis of a relatively high ED50 value for the inhibition of MF synthesis in vitro [121]. An in vivo bioassay was also established to measure MF titres in haemolymph samples of H. americanus and C. pagurus following the injection of putative MOIHs. Intriguingly, such bioassays identified active peptide fractions that are distinct from the previously identified and sequenced MOIHs of C. pagurus [121]. The peptide sequences of these unique MOIHs have not been published yet but the results indicate that there may be two groups of MOIHs acting on crustaceans: those having a direct effect on the MO (e.g. the MOIHs identified in in vitro assays), and those acting indirectly on the MO to elevate MF titres in the haemolymph. Should both groups of peptides be considered as physiologically relevant regulators of the MO when only one group demonstrates an effect in vivol Like JH III in insects, MF seems to play a role in enhancing crustacean reproductive maturity, as well as maintaining the juvenile morphology [123]. The synthesis of JH III in insects is also regulated by neuropeptides, one group of regulators (the allatostatins) is, in fact, similar in structure to neuropeptides that have been isolated from crustaceans (see Section 3.3) but there is no structural resemblance between the allatostatins and the MOIHs and there is no report on the role of allatostatins in regulating the crustacean MOs. Investigations into the mode of action of MOIH (or SG extracts) on the MO have been carried out in a few crustaceans only and not much is known. Cyclic nucleotides are involved in signal transduction - cyclic GMP is apparently important in H. americanus, whereas cyclic AMP is essential in C, pagurus [see 121].

102 102

3.2.5. Ion-transporting peptide Peptides that are structurally related to the cHH family of peptides are not restricted to crustaceans, although for a long time this was thought to be the case. In several insect species a peptide with structural features characteristic of the cHH has been isolated or deduced from cDNA sequences. This peptide is named "ion-transporting peptide" (ITP) because it stimulates the reabsorption of ions and water from primary urine in the hindgut of locusts [124], thus, it plays a role in osmoregulation. ITP was first isolated from CCs of S. gregaria and partially sequenced (amino acids 1-33); in a conspecific in vitro bioassay 1 -5 nM ITP stimulated reabsorption of Cl", Na+, K+ and fluid in isolated ilea [124, 125]. The complete peptide sequence of ITP from S. gregaria was later deduced from cDNA from locust brains; the 72 amino acid peptide (Table 3) has a free N-terminus, amidated C-terminus and the characteristic 6 Cys residues with about 40 % overall identity to cHHs from decapod crustaceans [126]. The in vitro bioassay used to identify ITPs, measured the active transport of Cl" from the lumen side of ileum preparations (mounted as a flat sheet), this transport is reflected by the positive change in short circuit current (Isc) across the ileum membrane [127]; compounds to be tested in this assay were always applied to the haemocoel side of the ileum preparation. In similar bioassays, crude extracts of CCs from a variety of orthopteran insects (migratory locust, crickets, cockroaches, stick insect) stimulated Cl" transport across the ileum of S. gregaria, whereas CC extracts from a lepidopteran and a dipteran showed no ITP activity in this heterologous assay, regardless of CC dosage applied [128]. This is most likely due to group-specific structural features of the peptide or its receptor, since peptides that are structurally homologous to the locust ITP are present in Lepidoptera and Diptera and further, cHH-family peptides from crustaceans have also not shown ITP activity in the locust ileal bioassay [129]. Putative ITP sequences (Table 3) have been deduced from cDNA of L. migratoria [128] and B. mori [130], as well as postulated from the genome of D. melanogaster [131]. In addition to these putative ITP sequences, an ITPlike peptide was identified, cloned and sequenced from a locust ileal mRNA library [126]. The ITP-like cDNA sequence was identical to that of the brain ITP cDNA except for an insert of 121 bp that preceded the

103 103

ITP C-terminus, and the predicted absence of a terminal amidation; that means, only the first 40 amino acid residues of ITP and the ITP-like peptide are identical [126]. This partial identity is reminiscent of the situation with C. maenas SG-cHH and PO-cHH and suggests also that the peptides arise from alternative splicing of a single gene (see Section 3.2.1). Like the crab PO-cHH [90], the non-amidated ITP-like peptide of the desert locust does not display the classic function in the original bioassay: instead of stimulating the ion/fluid transport in the ileal bioassay, recombinant ITP-like peptide inhibited the stimulation brought on by synthetic ITP [126]. Structure-function studies were conducted with recombinant ITPs that were suitably modified by site-directed mutagenesis. These studies are discussed in a recent review [129]. Briefly, the C-terminus is important for biological activity and Cterminally truncated ITPs, or the absence of the amide abolishes stimulation of ileal Isc; the 3 disulfide bridges are also important for activity. Interestingly, single deletions indicated that He 1 is important for receptor binding: when this residue is deleted in ITP-like peptides, the antagonistic effect to ITP is completely abolished. Hence, the observed inhibitory effect of ITP-like peptide on ileum transport appears to simply be a consequence of competitive receptor binding, with ITP-like peptide being able to bind to the ITP receptor but not being able to activate the receptor due to its changed C-terminus [129]. Factors from the eyestalks of H. americanus, specifically cHH, have been implicated in playing a role in osmoregulation too [132]. The involvement of cHH in osmoregulation was confirmed in another decapod crustacean, viz in the crayfish Astacus leptodactylus, where the injection of cHH into eyestalk-ablated specimens caused a significant increase in the haemolymph osmolality and Na+ concentration, whereas the concentration of Cl" remained unchanged [133]. 3.2.6. Sub-grouping of the cHH peptide family Members of this extensive and functionally diverse family of peptides that occur in arthropods show strong structural homology to each other, yet they can be sub-grouped on the basis of primary structure, as well as on preprohormone structure. Computer analyses, based on structural motifs of the mature peptides and their preprohormones, were carried out [134]. Thirty-two sequences were included in the analyses which showed that

104

the cHH peptide family subdivides into two groups, Group I contains the cHHs (including the MOIH from the spider crab, the MIH of P. bouvieri and the insect ITPs), while Group II contains the lobster VIH, MIHs and the remaining MOIHs. The preprohormone structure of Group I peptides is characterized by a signal peptide, a so-called cHH precursor-related peptide (CPRP), followed by a dibasic cleavage site and the sequence for the mature cHH (or ITP) sequence with an amidation signal at the Cterminus (see for example [135]). Preprohormones of Group II peptides are arranged into a signal peptide and the sequence for the mature VIH, MIH or MOIH peptide [134]. Many questions are still unanswered about the cHH peptide family: (a) there is clearly structural homology amongst the CPRPs [see 136] but what is the role of the CPRPs, if any? (b) how does the cHH effect its multi-functionality, while the VIH, MIH and MOIH seem to be more limited in functional activity, and is there any significance in this? (c) why are certain peptide functions not group- or species-specific and others are, and what does this imply about receptorligand interactions?

3.3. The allatostatin superfamily Two groups of hormones regulate development and reproduction in insects, namely the ecdysteroids and the JHs. In crustaceans too, ecdysteroids are involved in the hormonal control of growth, and a chemical compound, which is similar to the JHs of insects, is present and thought to play a role in crustacean reproduction and development. The insect JH is a species-specific acyclic sesquiterpenoid epoxide, which is synthesized in a pair of retrocerebral epithelial organs called the corpora allata (CA; see Fig. 1). In decapod crustaceans, MF is the unepoxidated form of the insect JH III and it is synthesized and secreted from the MOs (see Section 3.2.4). In insects and in crustaceans, the synthesis of both ecdysteroids and JH/MF are subject to control by neuropeptides. Although ecdysteroids are considered to be "growth promoting hormones" in both insects and crustaceans, these steroid hormones are differently regulated in the two taxa. In insects, the synthesis and release of ecdysteroids from the prothoracic gland is positively regulated by the prothoracicotropic hormone (PTTH) which is synthesized in neurosecretory cells of the brain and released from the CA [37]. PTTH was first completely identified in B.

105

mori as a 109 amino acid long polypeptide with an N-glycosylation site and 7 Cys residues which form disulfide bridges [10]. hi crustaceans, on the other hand, the synthesis of ecdysteroids in the YO is inhibited by MIH (see Section 3.2.2). There is no structural resemblance at all between PTTH and MIH. JH in insects maintains larval and nymphal characteristics during development and suppresses metamorphosis into the adult form; similarly, MF in crustaceans retards metamorphosis and larval development [123]. Two types of neuropeptides control the production of JH in the CAs of insects in vitro: (a) allatotropins, which stimulate the biosynthesis of JH and have only been found in insects to date, and (b) allatostatins (ASTs) which, inter alia, inhibit the biosynthesis of JH but which also have numerous other effects (mainly myoinhibitory) [137]. Neuropeptides that are structurally similar to the ASTs are also present in crustaceans and, therefore, we will review here what is known from the ASTs of insects and crustaceans. hi insects, one distinguishes between ASTs of three major structural forms [53, 137]: the "cockroach" ASTs or A-type ASTs which are characterized by the C-terminal pentapeptide sequence Y/FXFGLamide, the "cricket" ASTs or B-type ASTs which are generally characterized by Trp residues at position 2 and 9, and the "moth" ASTs or C-type ASTs which are only found in a few species to date (various moths and in D. melanogaster) and are 15 amino acid residues long with a pGlu residue at the N-terminus, a free C-terminus and two Cys residues forming an intramolecular disulfide bridge. Thusfar, only peptides of the "cockroach" AST-type have been found in crustaceans [138-140], and, hence, we will limit our inspection only to this group of ASTs which is common to insects and crustaceans. To date, more than 60 members of the A-type ASTs are structurally known from insects and about 20 species have been analyzed. Although only three species of decapod crustaceans were investigated, more than 60 isoforms are known in these crustaceans (see Table 4); interestingly, none is identical to the ASTs identified so far in insects [139]. Thusfar, in each insect and crustacean species investigated to date, multiple isoforms of the A-type Asts have been isolated or demonstrated to be present (see Table 4). Whereas all members of the A-type Ast family contain the relatively conserved pentapeptide core C-terminal of Y/FXFGLamide (for Leu there can be conservative changes such as He or Val but also Met in the blowfly, Calliphora vomitoria), the N-terminal part of the peptide can

106

be of variable length and may be blocked at the N-terminus. Some ASTs are only pentapeptides, while the longest ASTs have 30 residues (Table 4). Table 4. Primary structures of members of the allatostatiD family of peptides. Peptide name, code name Crustaceans Carma-AST 1 Carma-AST 2 Carma-AST 3 Carma-AST 4 Carma-AST 5 Carma-AST 6 Carma-AST 7 Carma-AST 8 Carma-AST 9 Carma-AST 10 Carma-AST 11 Carma-AST 12 Carma-AST 13 Carma-AST 14 Carma-AST 15 Carma-AST 16 Carma-AST 17 Carma-AST 18 Carma-AST 19 Carma-AST 20 Penmo-AST 1 Penmo-AST 2 Penmo-AST 3 Penmo-AST 4 Penmo-AST 5 Penmo-AST 7 Penmo-AST 8 Penmo-AST 9 Penmo-AST 10 Penmo-AST 11 Penmo-AST 12 Penmo-AST 14 Penmo-AST 15 Penmo-AST 16 Penmo-AST 17 Penmo-AST 18 Penmo-AST 19 Penmo-AST 20 Penmo-AST 21 Penmo-AST 22 Penmo-AST 23 Penmo-AST 24 Penmo-AST 25 Penmo-AST 26 Penmo-AST 27 Penmo-AST 28

Species

Sequence

Carcinus maenas, Penaeus monodon YAFGL-NH2 EAYAFGL-NH 2 C. maenas EPYAFGL-NHa C. maenas DPYAFGL-NH2 C. maenas NPYAFGL-NH 2 C. maenas SPYAFGL-HH 2 C. maenas ASPYAFGL-NH 2 C. maenas C. maenas, Orconectes limosus, P. monodon AGPYAFGL-NH2 GGPYAFGL-NH 2 C. maenas APQPYAFGL-NH 2 C. maenas ATGQYAFGL-NH 2 C. maenas PDMYAFGL-NH2 C. maenas EYDDMYTEKRPKVYAFGL-NH2 C. maenas YSFGL-NH2 C. maenas, P, monodon AGPYSFGL-NH 2 C. maenas GGPYSYGL-NHa C. maenas SGQYSFGL-NH 2 C. maenas SDMYSFGL-NH 2 C. maenas APTDMYSFGL-NH2 C. maenas C. maenas

P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon P. monodon

GYEDEDEDRPFYALGLGKRPRTYSFGL-NH2 MJEDEDAASLF&FGL-NH 2 PDAEESHKRDRLYAFGL-NH 2 DRLYAFGL-NHa TGGPYAFGL-NH 2 SAGPYAFGL-NH 2 SGHYAFGL-NH 2 ANQYAFGL-NH2 AGQYAFGL-NH2 TPSYAFGL-NH 2 PQRDYAFGL-NH 2 SDYAFGL-NH 2 ANQYTFGL-NH 2 ASQYTFGL-NH 2 SQYTFGL-NH 2 YTFGL-NH 2 SGHYNFGL-NHj GHYNFGL-NH 2 AGPYEFGL-NH 2 GGPYEFGL-NH2 AAPYEFGL-NH 2 GPYEFGL-NH2 SPYEFGL-NH 2 NPYEFGL-NH 2 NEVPDPETERNSYDFGL-NH 2 EVPDPETERNSYDFGL-NH 2 PETERNSYDFGL-NH 2

Reference

[138,139] [138] [138] [138] [138] [138] [138] [138-140] [138] [138] [138] [138] [138] [138, 139] [138] [138] [138] [138] [138] [138] [139] [139] [139] [139] [B9] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139]

107 Penmo-AST 29 Penmo-AST 30 Penmo-AST 31 Penmo-AST 32 Penmo-AST 33 Penmo-AST 34 Penmo-AST 36 Penmo-AST 37 Penmo-AST 38 Penmo-AST 39 Penmo-AST 40

P, P. P. P. P. P. P. P. P. P. P.

monodon monodon monodon monodon monodon monodon monodon monodon monodon monodon monodon

Orcli-AST 1 Orcli-AST 2

Orconectes limosus O. limosus

NSYDFGL-NH 2 YDFGL-NHa AGHYSFGL-NH 2 DRTYSFGL-NH 2 PSAYSFGL-NH2 pENMYSFGL-NH 2 DARGALDLDQSPAYASDLGKRIGSAYSFGL-NH 2 TARGALDLDQSPAYASDLGKRIGSAYSFGL-NH 2 SVAYGFGL-NH 2 TVAYGFGL-NH 2 (X)GIYGFGL-NH 2

[139] [139] [139] [139] [139] [139] [139] [139] [139] [139] [139]

SAGPYAFGL-NH 2 PRVYGFGL-NH 2

[140] [140]

SRPFGFGL-NH2

[142]

AGMYSFGL-NH2 GEGRMYSFGL-NH 2 PNYERMAGSRFNFGL-NH2 GPDHRFAFGL-NH 2 SLHYGFGI-NH 2 PYSFGL-NH 2 VPMYDFGI-NH 2

[142] [142] [142] [142] [142] [142] [142]

GRQYSFGL-NH a ADGRTYAFGL-NHz IPMYDFGL-NH2 TSSLYSFGL-NHs

[257] [257] [257] [257]

TTRPQPFNFGL-NH 2

[258]

Insects Grybi-AST-A6

Gryllus bimaculatus

Grybi-AST-A7 Grybi-AST-A9 Giybi-AST-AlO Gtybi-AST-All Grybi-AST-A12 Giybi-AST-A13 Grybi-AST-A14

G. G. G. G. G, G. G,

Carmo-AST-Al Carmo-AST-A2 Carmo-AST-A4 Carmo-AST-A5

Carausius morosus C. morosus C, morosus C. morosus

Drome-AST 4

D. melanogaster

bimaculatus bimaculatus bimaculatus bimaculatus bimaculatus bimaculatus bimaculatus

A complete list of sequences up to 1999 is available for insect members of this family [10,149],

The AST precursor, known only from insects to date, is characterized by a long leader peptide which consists of a signal peptide and another peptide of no known function, then follow clusters of ASTs interspaced by "acidic spacers", i.e. peptides with a high number of Asp and Glu residues and a resultant low isoelectric point [141], and a short tail peptide of unknown function [142]. When one considers the array of insect ASTs from an evolutionary view, it appears that the evolutionary more ancient insects, such as cockroaches and crickets, have a larger number of AST isoforms encoded on the precursor than more recent insect taxa, and it is speculated that this may be linked to functional diversity of the peptides: the ASTs of "older" insects have true allatostatic action in addition to other functions (see below), and those of the evolutionary "younger" insects have lost the allatostatic function [142]. For some insect species it is known that virtually all the encoded AST peptides of the preprohormone are, indeed, expressed. This was done exemplarily for the American cockroach by the employment of sensitive methods of mass spectrometry [143] showing additionally that a

108 108

particular internal dibasic cleavage site that is present in a peptide called Peram-AST-2 in the preproallatostatin is used to produce a small octapeptide. Such an octapeptide is quite ubiquitous and occurs not only in cockroaches, but locusts, moths and blowflies suggesting a possibly similar mechanism of prohormone processing. Is there a biological need or relevance for so many structurally similar peptides, such as the allatostatins, in one organism? In C. vomitoria different AST peptides have myoinhibitory actions on different segments of the gut, thus, different ASTs may have different targets within the same animal, and there is evidence of the insect ASTs having different potencies of action on one target [144]. Although direct radioligand-binding assays were successfully used to partially characterize an insect AST receptor [145], all the AST receptor sequences known to date are derived from molecular biological approaches. A G protein-coupled receptor was cloned from D. melanogaster and expressed in frog oocytes for a functional test; the results demonstrated that an A-type AST is the ligand for this receptor [16]. Structurally, the transmembrane regions of this AST receptor are related to the mammalian somatostatin/galanin/opioid receptors; in fact, most closely related to the galanin receptors although the peptides, galanin and AST, have little structural commonality. After the first AST receptor was cloned, similar receptors were cloned from P. americana, C. morosus, B. mori and a second one from D. melanogaster [137]. In a functional electrophysiological test, the characteristic C-terminal pentapeptide core sequence of A-type ASTs is able to mediate activation of the D. melanogaster [146] and P. americana receptor (G. Gade, H. Marco, R. Weaver and D. Richter, unpublished results). The latter study also showed that all 14 A-type ASTs from P. americana are potent in the electrophysiological functional assay, thus, it appears that there is no need for each peptide binding to a specific receptor protein. This contrasts with a report on a putative AST receptor from the cockroach Diploptera punctata, where a competitive binding assay showed that some of the ASTs have a higher affinity to the receptor [145]; the receptor has not yet been sequenced. Although present in a large number of insect species, the A-type ASTs apparently display an allatostatic effect (inhibiting the biosynthesis of JH in vitro) only in cockroaches and crickets. In other insect species, the conspecific AST peptides do not have an inhibitory effect on the CA in vitro [137, 147]. Moreover, immunocytochemical methods have identified numerous cells in a wide variety of insects that

109

react positively to A-AST antisera. Thus, not only cells in the brain but in most parts of the CNS, as well as in nerves innervating visceral muscles, midgut cells and haemocytes are immunopositive, and this distribution suggests, of course, different and multiple functions, inter alia also functions of A-ASTs as neurotransmitters and/or neuromodulators [137, 148]. To date, the following effects have been demonstrated by conspecific A-type ASTs in insects [137,149]: 1. Myoinhibitory actions of ASTs were shown in a number of insect species, for example, both spontaneous and proctolin-induced contractions of the Mndgut of D. punctata were inhibited; as were the peristaltic contractions of the ileum of C. vomitoria, the spontaneous contraction of the muscles of the oviduct of S. gregaria, the contraction of muscles from oesophagus and crop of the codling moth, Cydia pomonella, and the contraction of heart muscle in the German cockroach, Blattella germanica. 2. In B. germanica an endogenous AST inhibited the production of vitellogenin in vitro. 3. In D. punctata an endogenous AST also had a stimulating action: using ligatured midgets in vitro it was found that this AST stimulated the activity of the carbohydrases, amylase and invertase. Despite the similarity of the crustacean MF, in structure and function, to the insect JH, the synthesis of MF in the MO is probably not regulated by the numerous allatostatin peptides identified in crustaceans. Instead, the synthesis of MF is inhibited by MOIH (see Section 3.2.4). It should be noted, however, that the crustacean ASTs have not yet been fully tested for a role in the regulation of MO activity, so they could perhaps act on the MO together with the MOIHs. The crustacean AST peptides show a widespread distribution in immunocytochemical investigations with immunoreactive neurons localized in the brain, ventral nerve cord and commissural organs [139], the stomatogastric nervous system, entire CNS and the POs [140] and with immunoreactive fibres projecting into the walking legs, eyestalks (medulla extema, medulla interna and medulla terminalis) and into neurohaemal release sites, such as the suboesophageal ganglia, POs and the thoracic ganglia [140]. Such a distribution has been interpreted as clues that the crustacean ASTs play a role in neurotransmission or neuromodulation. In conspecific bioassays, the application of ASTs to hindgut preparations of crayfish in vitro resulted in a decrease in amplitude and

110

frequency of myogenic contractions; electrophysiological measurements also revealed a decrease in the cycling frequency of the pyloric motor rhythms which is generated by the stomatogastric ganglion. Since no AST immunoreactivity was found associated with the hindgut or its innervation, it is possible that the effect of ASTs on hindgut is not neuromodulatory [140]. A number of different crayfish species have now been shown to contain motor neurons in their stomatogastric nervous system that display AST-like immunoreactivity [150]. 3.4. Muscle activity regulated by various neuropeptide families An overwhelming number of fully characterized neuropeptides, especially in insects, have myotropic activity: one way or the other these peptides regulate the contractile activity of visceral and/or skeletal muscles. One possible reason for this abundance of sequenced neuropeptides that regulate muscle activity may be the successful usage of a very simple bioassay technique [see 151]. 3.4.1. Proctolin The first insect neuropeptide to be isolated and structurally identified from the American cockroach was the pentapeptide proctolin (RYLPT) [152]. It stimulates and modulates the contractions in visceral and skeletal muscles and has a role as co-transmitter but not as a neurohormone. The identical structure has been found in crustaceans (see Table 5): first in the PO of Homarus vulgaris [153] and later confirmed from the PO of C. maenas [154] here it has inotropic effects on the heart, i.e. it enhances the frequency of the spontaneous contractions. In the crab, C. borealis, proctolin has modulatory functions on neural circuits of the stomatogastric system [155]. More information, especially on structureactivity studies, breakdown and localization of proctolin-immunoreactive neurons can be gleaned from [10, 73, 156]. Based on binding studies, a receptor for proctolin was proposed in the locust oviduct [157]. Recently, with the help of the D. melanogaster genomic data base [158] and molecular cloning, a G protein-coupled receptor was identified, which, when expressed in HEK (human embryonic kidney) cells responded very sensitively (EC50: 3 x 10"10 M) to proctolin in a functional bioassay

111 Ill

measuring Ca2+ mobilization [159]. Another group found similar results and reported an EC50 value of 6 x 10"10 M [160]. It was also shown that the recombinant receptor binds proctolin in a competitive radioreceptor assay with high affinity (IC50 of about 4 x 10'9M) [159]. In addition, with the use of an antibody directed against the receptor, the latter authors demonstrated receptor-positive tissues in D. melanogaster which correlate well with those tissues/organs known to respond to proctolin. Interestingly, the genome of A. gambiae apparently does not contain an orthologue gene to the D. melanogaster proctolin receptor [159-161]. Previous studies in A. gambiae also reported an inability to detect proctolin-like immunoreactivity, proctolin-like biological activity, or proctolin responsiveness [159]. 3.4.2. Crustacean cardioactive peptide Another neuropeptide that has the same structure in crustaceans and insects is the cyclic nonapeptide, crustacean cardioactive peptide (CCAP; PFCNAFTGCamide) (Table 5), which was first isolated and sequenced from POs of C. maenas [162] and shown to have a stimulatory influence on the frequency of the heart beat in the shore crab and of the hindgut of O. limosus [163]. Later, the same peptide was isolated and sequenced from a number of insects such as migratory locust, tobacco hornmoth, southern armyworm (Spodoptera eridania), the mealworm Tenebrio molitor [10, 73] and the stick insect C. morosus [164]. Molecular biological and bioinformatic studies have identified or predicted genes which encode the precursor for CCAP in some insects, such as M. sexta [165], D. melanogaster [158, 166] and A. gambiae [70], but not in crustaceans. The CCAP precursor encodes a signal sequence of 20 amino acids followed by a dibasic cleavage site and a single copy of CCAP with a Gly residue (to form post-translationally the amidated peptide) and another two basic amino acid residues for proteolytic cleavage; no other putative peptides are encoded [165]. Localization of CCAP in crustaceans and various insect species has been studied in detail with the aid of immunocytochemistry [73, 167]. The distribution, for example, in neurons of brain, midgut and ventral nerve cord of insects suggests multiple and diverse functions, some of those are surely neuromodulatory (immunopositive interneurons which have processes into brain neuropil) but others are neurohormonal roles

112

(immunopositive processes to corpora cardiaca). It is not known what regulates the release of CCAP in crustaceans and insects, but in insects CCAP itself is a releasing factor for the well-known metabolic AKHs (see Section 3.1.1). Concentrations of CCAP were measured by RIA in haemolymph (10 pmol per liter) and POs (40 pmol per PO) of the shore crab [167]. CCAP is also released in massive amounts into the haemolymph of C. maenas and O. limosus just prior to ecdysis [168]. No detailed studies on inactivation of CCAP in any organism has been conducted. A G protein-coupled receptor for CCAP from D. melanogaster was cloned and functionally expressed in frog oocytes [59]. Although in this study the receptor also recognized the AKH of D. melanogaster to some extent, this can possibly be explained by some technical problems with the measurement as outlined elsewhere (169]. In the latter study CCAP appeared to be a very good candidate ligand for the cloned receptor with an EC50 of 5.4 x 10"10 M when expressed in CHO cells. The mode of action of CCAP is not known, but there are a large number of functions that are attributed to this peptide in crustaceans and insects: in crustaceans, it has an inotropic and a chronotropic (i.e. enhances the amplitude of spontaneous contractions) effect on the heart in the shore crab, a myotropic action on the hindgut of O. limosus, but no cardioactive effect in the American lobster and the Dungeness crab [167]. It has a neuromodulatory role or acts as neurotransmitter on the pattern of motor neurons that activate swimmeret beating in the crayfish P. leniusculus [170], Yet another effect of CCAP is the reduction of the amplitude of the electroretinogram in O. limosus [171]. In M potiuna high concentrations of CCAP were able to disperse the erythrophores; but this may rather be a pharmacological effect [63]. In insects, CCAP stimulates the contractions of heart, hindgut and oviduct in various species, but it is also one of the releasing factors for AKHs in the migratory locust [73, 172] and it is involved in inducing the motor programme that is so important for larval ecdysis and adult eclosion [73, 173]. In fact, a recent publication reports that genetic ablation of the CCAP neurons in D. melanogaster (although not lethal during the larval stages but lethal during pupal ecdysis) disrupts "the timing and organization of ecdysis behaviour [174].

113 Table 5. Primary structures of members of various peptide families with myoactivity. Peptide name, code-name

Species

Sequence

Reference

1. Proctolin Crustaceans Peram-proctolin

Homarus vulgaris and other crustacean species

RYLPT

[153]

Insects Peram-proctolin

Periplaneta americana and other insect species ;s

RYLPT

[152, 156]

PFCNAFTGC-NH 2

[162]

Carausius morosus

PFCNAFTGC-NH 2

[164]

Penaeus vannamei

ASFSPWG-NH 2

[177]

2. Crustacean cardioactive peptide Crustaceans Carma-CCAP Carcinus maenas Insects Carma-CCAP 3. (myo)kinins Crustaceans Penva-K-I Penva-K-II Penva-K-IH Penva-K-IV Penva-K-V Penva-K-VI Insects Peram-K-I Peram-K-II Peram-K-III Peram-K-IV Peram-K-V Peram-K-VI (= Locmi-KI) Peram-K-VII (= Leuma-K-VII) Peram-K-VHI (= Leuma-K-VIII) Musdo-K Drome-K.

P. vannamei

DFSAWA-NH2

[177]

P. P. P. P.

vannamei vannamei vannamei vannamei

PAFSPWG-NH 2 VAFSPWG-NH 2 pEAFSPWA-NH 2 AFSPWA-NH 2

[259] [259] [259] [259]

P. P. P. P. P. P.

americana americana americana americana americana americana

RPSFNSWG-NH 2 DASFSSWG-NH 2 DPSFNSWG-NH 2 GAQFSSWG-NH2 SPAFNSWG-NH 2 AFSSWG-NH 2

[260] [260] [260] [260] [260] [260]

P. americana

DPAFSSWG-NH 2

[260]

P. americana

GADFYSWG-NH2

[260]

NTVVLGKKQRFHSWG-NH2

[176, 261] [176, 180]

Musca domestica, Stomoxys calcitrans, Haematobia irritans D. melanogaster, Neobelliera bullata, Anopheles gambiae

NSVVLGKKQRFHSWG-NH2

4. FMRF-NH2-related peptides a) FMRF-NH? Crustaceans Do not have FMRF-NH2s Insects Neobu-FMRF

N. bullata

pEPSQDFMRF-NH2

[189]

SDRNFLRF-NH2 TNRNFLRF-NH2 GYNRSFLRF-NH2

[194] [194] [262]

b) FLRF-NHVmvosuppressins Crustaceans Homam-FLRF-I Homam-FLRF-II Calsa-FLRF

Homarus americanus H. americanus Callinectes sapidus

114 Procl-FLRF-I Procl-FLRF-II (=Macro-FLRF-I) Macro-FLRF-II Macro-FLRF-III Macro-FLRF-IV Macro-FLRF-V Macro-FLRF-VI Macro-FLRF-VII Macro-FLRF-VIII Insects Leuma-MS

Procambarus clarkii P. clarkii, Macrobrachium rosenbergii M. rosenbergii M. rosenbergii M. rosenbergii M. rosenbergii M. rosenbergii M. rosenbergii M. rosenbergii Leucophaea maderae, P. americana, Diploptera put

NRNFLRF-NH2 DRNFLRF-NH2 ADKNFLRF-NH2 NYDKNFLRF-NH2 APALRLRF-NH2 DRTPALRLRF-NH2 DGGRNFLRF-NH2 GYGDRNFLRF-NH2 VSHNNFLRF-NH2

[263] [263, 264] [264] [264] [264] [264] [195] [195] [195]

pEDVDHVFLRF-NH2

[190, 265, 266]

pEFDEY(SO3H)GHMRF-NH2

[197, 198] [197,

c) sulfakinins Crustaceans Penmo-SK-I Penmo-SK-II Penmo-SK-III Insects Peram-SK-I Blocked Peram-SK-I Glu-methylated Peram-SK-I Leuma-SK-II

P. monodon, P. vannamei Penaeus monodon, P. vannamei P. monodon

AGGSGGVGGEY*DDY(SO3H)GHLIRF-NH2

198] VGGEYDDY(SO3H)GHLRF-NH2

L. maderae, P. americana P. americana P. americana L. maderae, P. americana

EQFDDY (SO3H) GHMRF-NH2

[197] [267]

pEQFDDY(SO3H)GHMRF-NH2

[199]

(CH3) EQFDDY (SO3H) GHMRF-NH2 pESDDY (SO3H) GHMRF-NH2

[199] [268,

199] unblocked Leuma-SK-II

P. americana

QSDDY(SO3H)GHMRF-NH2

[199]

RARPRF-NH2 YSQVSRPRF-NH2 YAIAGRPRF-NH2 YSLRARPRF-NH2

[208] [208] [208] [208]

SNSRPPRKNDVNTMADAYKFLQDLDTYYGDR ARVRF-NH2 £SFTDARPQDDPTSVAEAIRLLQELETKHAQHA RPRPF-NH 2

[213]

d) NPFs Crustaceans Onlv short NPFs Penmo-PYF-I Penmo-PYF-II Penmo-PYF-III Penmo-PYF-IV

Penaeus monodon P. monodon P. monodon P. monodon

Insects Long NPFs Drome-NPF

D. melanogasler

Aedae-NPF

Aedes aegypti

Short NPFs Drome-NPF-I Drome-NPF-II Drome-NPF-III Drome-NPF-IV

D. D. D. D.

e) pvrokinins Crustaceans Penva-PK-I Penva-PK-II

P. vannamei P. vannamei

melanogasler melanogaster melanogaster melanogaster

[214]

AQRSPSLRLRF-NH2 WFGDVNQKPIRSPSLRLRPF-NH2 PQRLRW-NH2 PMRLRW-NH2

[166] [166] [166] [166]

DFAFSPRL-NH2 ADFAFNPRL-NHz

[219] [219]

115 Insects Peram-PK-I Peram-PK-II Peram-PK-III Peram-PK-IV Peram-PK-V Peram-PK-VI Perfu-PK-IV Carmo-PK Drome-PK-I Drome-PK-II

P. americana P. americana P. americana P. americana P. americana P. americana Periplaneta fuliginosa Carausius morosus D. melanogaster D. melanogaster

HTAGFIPRL-NH2 SPPFAPRL-NH 2 LVPFRPRL-NH 2 DHLPHDVYSPRL-NH 2 GGGGSGETSGMWFGPRL-NH2 SESEVPGMWFGPRL-NH 2 DHLSHDVYSPRL-NH 2 DEGGTQYTPRL-NH 2 TGPSASSGLWFGPRL-NH2 SVPFKPRL-NH 2

[269] [269] [270] [270] [270] [271] [271] [164] [272] [220, 221]

APSGFLGMR-NH 2 SGFLGMR-NH 2

[226] [226, 177]

APSGFLGVR-NH 2 APEESPKRAPSGFLGVR-NH 2 NGERAPGSKKAPSGFLGTR-NH 2 APSGFMGMR-NH2 APAMGFQGVR-NH2 APAAGFFGMR-NH2 VPASGFFGMR-NH 2 GPSMGFHGMR-NH2 APSMGFQGMR-NH2

[273] [273] [273] [273] [273] [274] [274] [274] [274]

f) tachvkinins Crustaceans Canbo-TRP-Ia CanboTRP-Ib

Cancer borealis C. borealis, P. vannamei

Insects Leuma-TRP-I Leuma- TRP-II Leuma- TRP-III Leuma- TRP-IV Leuma- TRP-V Leuma- TRP-VI Leuma- TRP-VII Leuma- TRP-VIII Leuma- TRP-IX

L. L. L. L. L. L. L. L L.

maderae maderae maderae maderae maderae maderae maderae maderae maderae

A complete list of sequences for insect peptides up to 1997 is available [10, 37]. Insect tachykinin-related peptides are given in [73]. *also sulfated in P. vannamei [198],

3.4.3. The (Myo)kinin family The (myo) kinin family of peptides is especially a very interesting one, because its members have two very different functional activities in insects. The first members of this family were originally isolated on the basis of stimulation of the cockroach hindgut in vitro from extracts of whole heads of L. maderae [151]. This species contains 8 isoforms of kinins. Shortly after their first isolation, it was reported that these kinins also stimulate diuretic activity of isolated Malpighian tubules of the yellow fever mosquito Aedes aegypti [175]. This function of kinins has now been shown regularly in conspecific diuretic assays with a number of other insect species. To date, the structure of kinins (see Table 5) have been fully elucidated in insects from the American cockroach (8 isoforms), the house cricket, Acheta domesticus (5 isoforms), the migratory locust (1 isoform), the mosquitoes, Culex salinarius and A.

116 116

aegypti (each 3 forms), housefly (M domestica), stablefly (Stomoxys calcitrans), hornfly (Haematobia irritans), flesh fly (Neobelliera bullata), D. melanogaster (each one form) and corn ear moth, Heliothis zea (3 forms) [129, 176]. In crustaceans, 6 kinin isoforms (Table 5) are known from the prawn Penaeus vannamei [177, 178]. Structurally, the kinin family is characterized by the C-terminal pentapeptide sequence PheXaa -Xaa -Trp-Gly amide; in a few crustacean kinins the ultimate Cterminal amino acid is however Ala (see Table 5). A preprohormone for kinins is reported from A. aegypti [179]. The organization is: a putative signal peptide (18 amino acids) followed by a 210 amino acid long prokinin which encodes one copy each of the 3 kinins of A. aegypti. The kinin precursor is also deduced from the genome of D. melanogaster [180], but no information has been gathered for any crustacean species. Localization of kinin-containing neurons has been achieved in many insect species by immunocytochemistry. Immunoreactivity with kinin antibodies occur in neurosecretory cells, in interneurons and in wellknown release sites but apparently not in neurons that directly innervate the hindgut and/or Malpighian tubules [10, 73]. Thus, it is suggested that kinins act as true neurohormones. RIAs have been developed and titre determinations executed but overestimation of kinin titres in non-purified haemolymph is possible; especially because the antisera are not well characterized and it is known that inactive breakdown products of kinins are recognized as well [see 129]. Degradation of kinins is well studied in the corn ear moth H. zea [181]: an enzyme that is similar to the vertebrate metalloprotease neprilysin is bound to membranes of the Malpighian tubules of the moth and cleaves an endogenous kinin primarily between Pro5 and Trp6, hence, destroying the C-terminal pentapeptide that is so important for biological activity. Structure-function studies had previously shown, when testing kinin analogues for their diuretic effects on cricket tubules in vitro, that this C-terminal pentapeptide is the minimum sequence required to elicit the same activity and potency as the parent molecule [10, 129]. Further studies revealed that the active conformation of the core sequence is a type VI P-turn where the aromatic side chains of the invariant Phe and Trp residues of the pentapeptide are very likely interacting with the receptor. The latter is G protein-coupled, has been cloned from D. melanogaster and expressed in heterologous cell lines and shown to interact well with the endogenous kinin of D. melanogaster [182]. The receptor is localized, as shown by

117

immunocytochemistry, in the secondary stellate cells of D. melanogaster, where these cells are the target for the kinin. Studies on its mode of action have demonstrated that signalling is dependent on activation of PLC, IP3 production and increase of intracellular Ca +, resulting in an activated Cl~ shunt which may be located transcellular through apical and basal Cl" channels [129, 183]. Apart from their role as true neurohormones in insects, the insect kinins may also fulfil neuromodulatory roles on specific neuronal circuits, as deduced from immunocytochemical studies with various insect species [see 73]. In crustaceans, the kinins were isolated first on the basis of a heterologous bioassay, namely to stimulate the contraction of the hindgut of the Madeira cockroach but the crustacean kinins were also active in another insect assay, i.e. they stimulated fluid secretion in the Malpighian tubules of A. domesticus [177]. Later, the 6 kinins from the prawn (Table 5) were tested in another heterologous bioassay, but at least it was on crustacean tissues: all prawn kinins were able to stimulate the basal tonus of the isolated oviduct of A. leptodactylus and also enhanced the frequency and amplitude of the spontaneous contractions of the isolated hindgut of this crayfish [184]. Conspecific assays on the hindgut of the prawn were only successful when the endogenous peptide Penva-K-1 was used at a concentration of 10"6 M [178]. 3.4.4. The FMRFamide-related peptide superfamily There is a very diverse superfamily of neuropeptides present in most invertebrates that are called FMRFamide-related peptides (FaRPs); the name is based on the first member of this group isolated from ganglia of a clam (mollusc) and identified as Phe-Met-Arg-Phe amide (FMRFamide); this peptide had a strong cardioexcitatory action [185]. A number of important families are grouped into this superfamily: the FMRFamides and extended FMRFamides, myosuppressins (FLRFamides), sulfakinins (HMRFamides) and, neuropeptides F and Y (NPFs and NPYs). We will not attempt to review all these peptides in detail but will make only a few comments and dwell a bit deeper into the sulfakinins, NPFs and NPYs as far as resemblance between insect and crustacean peptides is concerned.

118

3.4.4.1. The FMRFamides FMRFamides have been found in a variety of insects and detailed accounts of their distribution, structure, preprohormone organization and function have been given in previous reviews [10, 73, 186, 187]; Table 5 shows an example of a FMRFamide. Recently, a G protein-coupled receptor was cloned from D. melanogaster which, when stably expressed in CHO cells, reacted with high affinity (EC50: 9 x 10~10 M) to one of the 8 endogenous FMRFamides and with lower affinity to some others, suggesting that more FMRFamide receptors must exist in D. melanogaster [188]. The same receptor was also cloned by another research group [189] and their experiments did not show this differential affinity of the receptor for the endogenous peptides. Thus, there is a disagreement in the results of these two research groups with respect to the rank order potencies of the different ligands. 3.4.4.2. The FLRFamides A close structural relative of the FMRFamides are the FLRFamides. Its first candidate in insects was characterized as a myosuppressin from L. maderae where it inhibits the spontaneous contraction of the hindgut [151]. Thereafter, a number of FLRFamides were isolated from various insects (see Table 5 for post-1997 sequences) [73, 187]. The cockroach D. punctata contains the same myosuppressin as L. maderae in its brain. The cDNA encoding the precursor for that peptide was isolated and shown to have a single copy of the myosuppressin at its C-terminus and, additionally, the necessary sites for cleavage and amidation to generate the mature peptide [190]. A single-copy gene for a myosuppressin is also found in the genome of D. melanogaster [191]. Recently, two G proteincoupled receptors were identified in D. melanogaster, by cloning their genes and expressing the cDNA in CHO cells, these receptors were shown to be activated by low concentrations of the endogenous fly's myosuppressin (EC50: 4 x 10"8 M) [192]. These results were confirmed and expanded on in a more substantive study that made use of a fluorescence translocation assay to identify ligands of G protein-coupled receptors (Johnson, E.C.; Bohn, L.M.; Barak, L.S.; Birse, R.T.; Nassel,

119

D.R.; Caron, M.G.; Taghert, P.H.; personal communication). This assay is based on the rationale that most G protein-coupled receptors have a common mechanism to terminate the hormonal signal after receptor activation. In vertebrates, such receptors are desensitized by the action of G protein-coupled receptor kinases and |3arrestins. A previous report showed the desensitization of known and diverse mammalian G proteincoupled receptors with chimeras of parrestin2 tagged with green fluorescent protein (Parr2-GFP) [193]. In such an assay, unstimulated cells which expressed the receptor showed a diffuse distribution of GFP fluorescence in the cytoplasm and when these cells were exposed to the appropriate ligand (which binds the receptor), most of the fluorescence (i.e. Parr2-GFP) then translocated to the cell membrane. Using this Parr2GFP translocation assay, it was shown that 1 x 10"6 M Dromemyosuppressin triggered the translocation of Parr2-GFP to the cell membrane of HEK cells that expressed receptor cDNA of the D. melanogaster gene CG 13803, as well as in cells expressing the paralogous receptor gene CG 8985 (Johnson, E.C.; Bonn, L.M.; Barak, L.S.; Birse, R.T.; Nassel, D.R.; Caron, M.G.; Taghert, P.H.; personal communication). In accordance with its inhibitory physiological action, the activation of CG 13803 and CG 8985 receptors by Dromemyosuppressin, in the presence of forskolin, caused a decrease in the levels of cyclic AMP, hinting that the receptor couples via inhibitory G proteins (Johnson, E.C.; Bohn, L.M.; Barak, L.S.; Birse, R.T.; Nassel, D.R.; Caron, M.G.; Taghert, P.H.; personal communication). In crustaceans true FMRFamides are not known, but only FLRFamides (Table 5). They were first isolated from the PO of//, americanus [194] and later found in crabs, crayfish and a prawn [195]. Their functional role is most likely neurohormonal, i.e. probably cardioexcitatory and influencing contractions of visceral muscles, as well as neuromodulatory, i.e. modulating motor patterns in the stomatogastric ganglion [150]. 3.4.4.3. The sulfakinin family The first sulfakinins were again isolated from whole heads of L. maderae and its sequence elucidation suggested that this family is structurally related to the mammalian peptide family comprising gastrin and cholecystokinin [151]. Its biological activity was measured by stimulation of contractions of the cockroach hindgut. Later, members of this family

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were found in a number of insects, such as the American cockroach, the migratory locust, and various dipteran species, including D. melanogaster [37, 73] (see Table 5), In crustaceans, sulfakinins have been characterized only in the two prawn species P. vannamei [196] and P. monodon [197]. The characteristics of this family in insects and crustaceans are a Cterminal YGHMRFamide and a sulfated Tyr residue (Table 5). In the prawn P. vannamei, one of the two sulfakinins is even sulfated at both Tyr residues [198]. Non-sulfated forms apparently also occur in insects, hi the American cockroach, other post-translational modifications have taken place, namely a pGlu residue at the N-terminus or even Omethylation of the N-terminal glutamic acid [199] (see Table 5). The precursor of sulfakinins has been analyzed in D. melanogaster. it contains three putative peptides of which two are clearly sulfakinins [200]. A G protein-coupled receptor has been cloned from D. melanogaster; when it was functionally expressed in CHO cells, the endogenous Dromesulfakinin was the most active ligand [201]. Imrnunocytochemical studies revealed that sulfakinins are predominantly located in the CNS and are only found at neurosecretory release sites of a few insects; sulfakinins, apparently, act as central neuromodulators and do not have a neurohormonal role [73]. Structure-activity studies demonstrated that non-sulfated analogues were inactive and that the C-terminal hexapeptide is the smallest functioning fragment (possessing about 10 % of the myotropic activity of the parent molecule) but full potency and efficacy is achieved by the C-terminal octapeptide [see 10]. Whereas the C-terminal peptide is HMRFamide for the sulfakinins, it is WMDFamide for gastrin and cholecystokinin which are also both sulfated. Although the vertebrate peptides as such are not active in the cockroach hindgut assay, replacement of the Asp residue with an Arg residue transformed them into active analogues in this bioassay [202]. The functional tasks of insect sulfakinins, besides stimulation of the contractions of the hindgut, are stimulation of heart beat in cockroaches, induction of the secretion of aamylase in the midgut of a beetle and reduction of the amount of food eaten by locusts and cockroaches upon injection of sulfakinin [53, 73], Similar actions (induction of satiety, secretion and muscle contractions) are known from the mammalian counterpart, cholecystokinin [203]. The crustacean sulfakinins have only been proven to have an effect on the cockroach hindgut; they were not active at all on hindgut or oviduct of the crayfish [198].

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3.4.4.4. The NPY superfamily, including the insect NPF family The vertebrate NPY family contains such prominent members as pancreatic polypeptide (PP), characterized initially from chicken where PP is synthesized and released from the endocrine cells of the pancreatic islets [204], neuropeptide Y (NPY) found in porcine brain [205] and peptide YY (PYY) isolated from porcine intestine [206]. Such peptides are characterized by a chain length of 36 amino acids and Tyr-amide at the C-terminus. Whereas NPY is exclusively expressed in neurons of mammals and is known to control processes such as stimulation of food intake, vasoconstriction, sexual behaviour and circadian rhythm, PP and PP Y are synthesized in endocrine cells of the gut and are inhibitors of gut motility and of the secretion of exocrine products from the pancreas. In insects, members of the NPY superfamily have been identified, but here they are called members of the neuropeptide F (NPF) family because their C-terminus is characterized by a Phe-amide (Table 5). In fact, NPFs have been elucidated before in a number of other invertebrate taxa such as Cestoda (tapeworms) and Mollusca [see 73]. According to chain length, insects possess long NPF-like peptides and short ones (see Table 5 for some examples). This distinction appears to be justified not only because of the different chain length but also because in D. melanogaster, different genes have been identified to code for short and long NPF-like peptides; similarly, the two types seem to have different receptors [73]. Fully characterized short NPF-like peptide structures are known from the insects A. aegypti, Leptinotarsa decemlineata, P. americana [207], H. zea, S. gregaria and are predicted for D. melanogaster and A. gambiae [70,73,158]. The C-termini of these peptides are either PXLR/KL/TRFamide or RPRFamide. In the prawn, P. monodon, four short NPF-like peptides have been isolated from the eyestalks and the primary sequence determined (Table 5); the one hexapeptide and three nonapeptides share the four C-terminal amino acids RPRFamide [208]. For the so-called A. aegypti "head peptide" the encoding gene has been cloned and the preprohormone is characterized by coding for a signal peptide, three identical "head peptides" and a 38-mer at the carboxy terminus that does not resemble any known insect neuropeptide [209]. A receptor that can be activated by the D. melanogaster short NPFs was recently identified [210, 211]. Expression of the receptor in CHO cells [210] or X. laevis oocytes [211] results in some different data for the

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potencies of the endogenous D. melanogaster NPFs; whereas they were equipotent in CHO cells, the electrophysiological results with the oocytes point to a higher potency for Drome-NPF 1. The receptor is expressed in brain, gut, Malpighian tubules, fat body of D. melanogaster larvae, as well as in ovaries of adult females [210]. A variety of functional activities have been attributed to short NPFs of insects, ranging from activation of muscle activity and inhibition of host seeking behaviour (A aegypti "head peptide") to stimulation of ovarian growth and an increase of vitellogenin titres in the haemolymph, thus myotropic and gonadotropic effects are suggested [73,209]. It is not known, however, whether such gonadotropic effects are direct or indirect, i.e. via stimulation on the ecdysone or juvenile hormone systems. In D. melanogaster the endogenous NPF is reported to control foraging and social behaviour [212]. In brief, the level of NPF RNA was high in the CNS tissues of larvae attracted to food, gene expression is, however, turned off in those larvae that showed aversion to feeding. When the feeding response was tested in transgenic larvae, in which the NPF neurons were ablated, it resulted in a premature insensitivity to the feeding stimulus. Moreover, developmental downregulation of NPF also suppresses cooperative burrowing behaviours which are normally displayed by older larvae; thus, the NPF system is necessary for social interaction as well [212], Immunocytochemical studies on the distribution of these peptides have not been very helpful and are ambiguous, largely because no specific antisera have been developed (the antisera would most likely cross-react with other members of the FaRFamide superfamily). Long NPFs have only been isolated in insects but not in crustaceans (see Table 5). The insect forms are known from D. melanogaster [213] and A. aegypti [214] and an orthologue has been found in the genome of A. gambiae [70]. The prepropeptide is organized into signal peptide and the 36 amino acid long NPF peptide and this organization is common to all members of the NPF superfamily of this length [213]. The long NPFs are found in brain neurosecretory cells and midgut endocrine cells as shown by imrnunocytochemistry and Northern blots [214]. Some years ago, a putative receptor that was structurally similar to the vertebrate NPY receptors had been cloned from D. melanogaster, first called PR4 and later re-named NepYr [215, 216]; recent data has questioned its role as a true receptor for the long NPF of D. melanogaster [217]. hi the latter study a novel receptor was cloned, called DmNPFRl, which was stably expressed in CHO-K1 cells along with the NepYr. Only DmNPFRl-

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expressing cells showed high affinity binding of the endogenous NPF in a radioreceptor assay and inhibition of forskolin-stimulated activity of AC; NepYr-transfected cells were negative in both assays [217]. The receptor is localized in the CNS and midgut of larvae of D. melanogaster. Genome analysis of A. gambiae has identified an orthologue of the DmNPFRl gene [161]. As to the function of long NPFs in insects, there is no conclusive evidence but it is speculated, mainly on the basis of its distribution, that it plays a role in regulation of feeding behaviour and digestion, thus acting very likely in a neuromodulatory capacity and as true hormone [213]. 3.4.5. The pyrokinin family and structurally related peptides Peptides of this family have received their name from the first pyrokinin (PK; isolated from L. maderae in which it was demonstrated to have a stimulatory action on the hindgut), which is characterized by a pGlu residue at the N-terminus and the pentapeptide F/YXPRLamide at the Cterminus [10, 73]. In other insects, peptides with an identical C-terminus to that of the PKs have been found and have been named myotropins (MTs), pheromone biosynthesis-activating neuropeptides (PBANs), diapause hormones (DHs), puparium acceleration factor, and melanization and reddish colouration hormones (MRCHs), according to the biological effects these peptides have been attributed to [10, 73]. PKs sensu strictu have been identified in cockroaches, locusts, and D. melanogaster [10, 73]. In P. americana, for example, six PKs are present (Table 5): whereas the isoforms 1 to 4 and 6 are produced in the neurosecretory cells of the suboesophageal ganglion and tritocerebrum, isoform 5 (and much less so isoform 6) is the main candidate present in the abdominal neurosecretory cells of the perisympathetic organs [73, 218]. In crustaceans, two PKs have been identified from P vannamei (Table 5) [219]. In D. melanogaster there are different genes encoding putative peptides of this family. One is called capability and encodes, in addition to two cardioacceleratory peptides with the C-terminal FPRXamide (cap 2b-1 and cap 2b-2), a third peptide (cap 2b-3) with the C-terminal FXPRL characteristic of the PK family [191]. The second gene is called hugin and encodes two peptides, one of which belongs to the PK family (Drome-PK-2); the other peptide called hug y, is rather related to the family of ecdysis-triggering hormones (ETHs) which lack

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the characteristic Phe residue in the C-terminus of PKs [220, 221]. Genes and precursors of PBANs/DHs have also been cloned from various Lepidoptera [73]. A G protein-coupled receptor was cloned and expressed in frog oocytes and shown to be most sensitive to Drome-PK-2 (and hug y) and had moderate sensitivity to cap 2b-3 and ETH [59]. Not surprisingly, considering the variety of peptides belonging to the PK family, there is also an array of functions known: the "true" PKs/MTs are myostimulatory in action on foregut, hindgut, oviduct and hyperneural muscle of locusts and/or cockroaches [73, 218], whereas the two crustacean PKs are potent stimulators of the hindgut of A. leptodactylus but are not active on the oviduct of this crayfish [184]. The other members of this peptide family are well known for their ability to stimulate the bioysnthesis of sex pheromones in a number of Lepidoptera (PBANs), to regulate melanization in larvae of some moths (MRCHs), to induce diapause in silkworm eggs (DHs), and to accelerate the formation of a puparium in flies [10, 73]. Extensive structure-activity studies have been performed, especially on PBAN. This peptide is also best analyzed with respect to: (a) analogues that display antagonistic effects, (b) pseudopeptides which have better penetration of the insect cuticle, and/or (c) resistance to peptidases [222, 223]. Such studies have been conducted in the context of pest control. The interested reader is referred to a recent overview which discusses some ideas centred around the possibility of using insect neuropeptides as agents to control pest insect species [53]. 3.4.6. Tachykinin-related peptides The best-known member of the vertebrate neuropeptide tachykinin (TK) family is the undecapeptide, substance P (RPKPQQFFGLMamide), which has diverse actions as excitatory neurotransmitter and also as modulator of various functions, including sensory processing, control of movement, gastric mobility, vasodilation and salination [10, 224]. The migratory locust was the first insect in which neuropeptides were found which had limited structural homology to vertebrate TKs; they are characterized by the C-terminal pentapeptide FX'GX2Ramide (see Table 5) instead of the FXGLMamide of vertebrate TKs [225]. To date, such peptides have been isolated and up to 9 isoforms per species sequenced from the migratory locust, the Madeira cockroach (Table 5), the blowfly C. vomitoria, the mosquito C. salinarius and the stable fly S. calcitrans

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[224]. In D. melanogaster, 6 TK-related peptides have been identified by molecular biological methods, including a slightly modified form [73] and in A. gambiae a gene has been identified which is predicted to encode TK-related peptides [70]. hi crustaceans, 2 TK-related peptides were found in C. horealis [226], one of which is also present in P. vannamei (Table 5) [177]. The precursor gene is only known from D. melanogaster. It consists of 4 exons, but only exon 2 and 3 encode multiple isoforms of TK-related peptides, viz. 4 isoforms on exon 2 and a further 2 forms on exon 3. Additionally, other peptides are encoded on this precursor but they are not related to known insect neuropeptide families, and no function is known either [227]. The first receptor (belonging to the G protein-coupled type) in insects that have some sequence similarities (in the transmembrane region) with mammalian TK receptors were cloned as early as the beginning of the 1990s from D. melanogaster [228, 229]. Whereas the earlier study used the mammalian substance P to activate the receptor, the later study tested, heterologously, the TK-related peptide II from L. migratoria when the receptor cDNA was stably expressed in mouse NIH-3T3 cell lines. Recently, expressing the cDNA of the 2 putative receptors for TKs from D. melanogaster and parr2-GFP cDNA in HEK-293 cells, it was demonstrated that both these paralogous receptors translocate the parr2-GFP to the cell membrane upon exposure to the putative Drome-TK-I [Johnson, E.C.; Bohn, L.M.; Barak, L.S.; Birse, R.T.; Nassel, D.R.; Caron, M.G.; Taghert, P.H.; personal communication]. Later, a similar receptor was cloned from 5. calcitrans [230] and, when expressed in a stable D. melanogaster Schneider 2 cell line, a number of insect TK-related peptides, including the conspecific one from the stable fly, were found to be potent agonists [231]. Moreover, structure-activity studies using the functional system described above, suggested that (1) the conserved residues Phe and Arg at the C-terminus are essential for receptor interaction, (2) the C-terminal pentapeptide FTGMRamide is the active core region of these peptides which display a large percentage of the possible maximal activity and (3) in accordance with its C-terminal amino acid, Arg-amide in insect TK-related peptides and Met-amide in mammalian TK, the respective insect and mammalian receptors show increased activity with their "own" type of C-terminal peptide [231]. In insects, TK-related peptides are mainly localized in brain and gut tissue. Their major actions are to stimulate title contractility of muscles in the cockroach hindgut and the locust oviduct, but it has also been shown

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in some insects that TK-related peptides stimulate muscles of fore- and midgut, the heart and even of Malpighian tubules; they also have a myomodulatory action on the extensor tibiae muscle of the migratory locust, are apparently involved in the control of release of adipokinetic hormone from the neurosecretory cells in the CC of this insect, and have been attributed various neuromodulatory roles in photoreceptors of the crayfish P. leniusculus and on motor patterns of the stomatogastrie ganglion of the crab C. borealis [73]. CONCLUSIONS: The analysis of the structures, function and modes of action of major neuropeptide families of insects and crustaceans has revealed the following: 1. a few peptides, such as proctolin and CCAP belong to a "one member family", they are ubiquitously distributed in invertebrates and occur in both taxa under review here; their function has apparently not changed during the course of evolution. 2. some peptides belong to families that are, to date at least, only known with certainty to occur in arthropods. These are the AKH/RPCH (in short, AKH) family and the eHH/MH/VIH/MOIH/ITP (in short, cHH) family. In both families, the members found in the crustaceans primarily have different functions to those found in insects. The differences between the two peptide families are clear: (a) most variant members of the AKH family have been sequenced from insects, whereas most members of the cHH family occur in crustaceans. (b) In the AKH family, only one member is found in crustaceans and this member is also present in at least one insect species. However, whereas it functions as a chromatophorotropin in crustaceans, it controls metabolism in the insect. The receptors in both groups are rather specific but will still react to other peptide members of the family: the crustacean RPCH and the insect AKHs cross-react biologically in the other's system. (c) In the cHH family there are (as yet) no identical members in any insect and crustacean, and the particular members do not cross-react in the other's system, i.e. ITP will not increase the glucose concentration in the circulation of a crustacean and the

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crustacean cHHs are not potent in affecting the transepithelial potential of a locust's hindgut. Some families have a number of members but are not restricted to arthropods, for example, the FaRPs which occur in most major taxa, including vertebrates. Their myoactive effect also appears to be ubiquitous. Although it is not always clear why numerous isoforms of a peptide family do exist in organisms (with respect to physiological relevance), it is interesting to see just how these multiple forms are produced in insects and crustaceans: (a) by distinct genes that may (or may not) also code for an unrelated peptide, e.g. the 2 cHH isoforms of the lobster, the 3 AKHs of the locust and the 2 MOIHs of the crab (b) by a single gene, e.g. all 14 allatostatins of the cockroach are encoded by one gene (c) by alternative splicing sites on one gene, e.g. ITP and ITP-like peptides in insects, and the PO-cHHs in crabs. The complete subject area of ligand/receptor co-evolution cannot really be answered and fully exploited since there is no detailed sequence information on receptors from crustaceans. The chief reason for this is because the complete genome of a crustacean has not been sequenced yet. Perhaps major progress in this area will be achieved when industry is convinced of the profitability of transgenic lobsters! ABBREVIATIONS PO CC XO SG CNS RPCH HPLC RIA EIA cHH ELISA PCR

= pericardial organ = corpora cardiaca = X-organ = sinus gland = central nervous system = red pigment-concentrating hormone = high pressure liquid chromatography = radioimmunoassay = enzyme immunoassay = crustacean hyperglycaemic hormone = enzyme-linked immunoassay = polymerase chain reaction

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AKH GIRK FAB MALDI ESI HrTH pGlu CHO PLC IP3 AC MO DRPH PDH PDF ORF PKA MIH YO PKG VIH GIH MOIH MF JH ITP Isc CA PTTH AST HEK cells CCAP FaRP NPF NPY Parr2-GFP PP PYY PK

= adipokinetic hormone = G protein-gated inwardly rectifying potassium channel = fast atom bombardment = matrix-assisted laser desorption/ionization = electrospray ionization = hypertrehalosaemic hormone = pyroglutamic acid = Chinese hamster ovary = phospholipase C = inositol trisphosphate = adenylate cyclase = mandibular organ = light-adapting distal retinal pigment hormone = pigment-dispersing hormone = pigment-dispersing factor = open reading frame = cyclic AMP-dependent protein kinase = moult-inhibiting hormone = Y-organ = cyclic GMP-dependent protein kinase = vitellogenesis-inhibiting hormone = gonad-inhibiting hormone = mandibular organ-inhibiting hormone = methyl farnesoate = juvenile hormone = ion-transporting peptide = short circuit current = corpora allata = prothoracicotropic hormone = allatostatin = human embryonic kidney cells = crustacean cardioactive peptide = FMRFamide-related peptide = neuropeptide F = neuropeptide Y = Parrestin2-green fluorescent protein = pancreatic polypeptide = peptide YY = pyrokinin

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MT PBAN DH MRCH ETH TK

= myotropin = pheromone biosynthesis-activating neuropeptide = diapause hormone = melanization and reddish colouration hormone = ecdysis-triggering hormone = tachykinin

ACKNOWLEDGEMENTS The authors thank Dr. L. Auerswald (University of Cape Town) for his help with the preparation of the manuscript and figures, Prof. P. Taghert (Washington University School of Medicine, St. Louis, USA) for critically reading some sections on myotropic peptides and The National Research Foundation (Pretoria, South Africa; grant number 205 3396) and the University of Cape Town (staff award to G.G.) for financial support.

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