Advances in Insect Physiology, Volume 29

Contributors S. Caveney D(7~m'lment ~/Zoo/ogy, University 0[ Western Ontario, Londml, Ontario N6A 5B7, Camula G. M. Coa...

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Contributors S. Caveney D(7~m'lment ~/Zoo/ogy, University 0[ Western Ontario, Londml, Ontario N6A 5B7, Camula G. M. Coast D~Tmrtment ~?/"Biology, Birkbeek ( Universitv ~/ London ), Malet Street, Lottdon WC1E 7tt)C UK B. C. Donly Southern Crop Protection am/Food Research Centre, A~rieulture and A~ri-f~md Canadtt, Lomlon. Ontario, Canada N5V 4T3 M. J. Hall Deparmtent qf Biological Sciences, The Open University, Milton Keynes, MK7 6AA, UK M. L. Hudson D~7~armlent o/ Biologie~tl Structure and Function, 611, S W Cam~ms Drive, SD, Ores.on Health and Sciences University, Port[and, OR 97201. USA D. B. M o r t o n

Del~arlment qf Biological Struetm'e and k)mction, 611. S W Campus Drive, SD, Ore,~on Health and Sciences University, Portland, OR 97201. USA I. Orchard

Universilv ¢?/' Toro/lto, Department ~?/ Zoology, 25 Harbor Street, Toro/tm, Ontario M5S 3G5. Can~ttkt J. E. Phillips Universil|' o/ British CohmdEa, DC/)~IPllIIClZl ()/ Zoolo£,yl', ~/alleozlvel', British Colmnhia V6 T 1Z4, Cana~kt D. J. Robinson D~7~artment ~?/ Biolo~zical Science.,,'. The Opetl Universill', ,]/[illoll Keynes, MK7 6AA. ("K D. A. Schooley DeparmTent ~?/Biochemistry ( 330 ) , Universi O, ~?/Nevada, N V 89557-0014. USA

Cyclic GMP Regulation and Function in Insects David B. Morton and Martin L. Hudson Department of Biological Structure and Function, 611, SW Campus Drive, SD, Oregon Health and Sciences University, Portland, OR 97201, USA

1 Introduction 2 2 Cyclic GMP regulation 2 2.1 Guanylyl cyclases 2 2.2 Phosphodiesterases 22 3 Cyclic GMP function 26 3.1 Molecular targets 26 3.2 Physiological functions 32 4 Concluding remarks 43 Acknowledgements 44 References 44

Abstract Guanosine 3'5' cyclic monophosphate (cGMP) mediates a wide variety of physiological processes in many invertebrate and vertebrate species. Here we discuss our current understanding of c G M P regulation and function in insects, reviewing components of the c G M P signaling cascade and some of the major physiological roles played by c G M P in insects. The recent completion of the Drosophih~ genome project has enabled us to identify all the potential elements of the c G M P signaling cascade in a single insect. Most of these proteins have not been previously characterized, and by comparing their predicted sequences with identified proteins from other species (insects and mammals) we describe their expected properties. The list of potential proteins that regulate c G M P levels includes five receptor guanylyl cyclases (GCs), two receptor-like GCs, five soluble GC subunits, a possible GC-activating peptide, five possible GCAP-like proteins and five phosphodiesterases that are predicted to hydrolyze cGMP. Downstream elements of c G M P signaling include two phosphodiesterases that could be regulated by cGMP, three cGMP-dependent protein kinases and two ion channels that could be regulated by cGMP. ,M)VANC'ES IN INSECT PHYSIOLO(IY ]SBN I) 12 I)24229 X

VOL

29

('~)l~Ir~i~llt ~ 2002 E/~uw~,r S~iemc L i d 41/~i~,IiI~ ql r~Tn'nduction m aHl I m m ~'~'~'p vud

2

D.B. MORTON AND M. L. HUDSON

1 Introduction The intracellular messenger, guanosine 3'5' cyclic monophosphate (cGMP) was first discovered shortly after the seminal work of Sutherland and Rall (1957) had established the concept of second messengers (Ashman et al., 1963). Since that time many studies in a wide variety of tissues and organisms have demonstrated the crucial role that c G M P plays in many physiological processes. While the majority of these studies have been in vertebrate preparations. there have also been several major contributions to c G M P research that have utilized insects. These include one of the first demonstrations of a cGMP-dependent protein kinase (PKG) (Kuo and Greengard, 1970), the first identification of nitric oxide (NO)-insensitive soluble guanylyl cyclases (GCs) (Nighorn el al., 1999; Simpson et al., 1999), and the demonstration that a naturally occurring behavioral polymorphism is due to mutations in a P K G gene (Osborne et al., 1997). This review describes the molecular components of the c G M P signaling system that have been identified in insects, and surveys the newly available genomic sequence of Drosophila to demonstrate the variety of genes that might be involved in the regulation and function of cGMP in a single organism. In addition, we review the signaling cascades of several important physiological processes in insects that are regulated by cGMP.

2

Cyclic GMP regulation

The intracellular concentration of c G M P in a cell is regulated by a balance between its rates of synthesis and breakdown. The synthesis of c G M P from GTP is catalyzed by GCs, and c G M P is hydrolyzed by phosphodiesterases (PDEs) to form GMP. There are several classes of both enzymes, and the activity of both is regulated by a wide variety of factors. Molecular cloning has identified several members of GCs and PDEs in insects, and the recent completion of the Drosophila Genome Project has now allowed a more complete description of the variety of possible regulatory mechanisms for controlling c G M P levels in insects. 2. l

GUANYLYLCYCLASES

Early studies investigating GC enzyme activity revealed that most mammalian tissues contained both cytoplasmic and membrane-bound GC activity (Tremblay et al., 1988). Although GC activity has been measured in a number of insect tissues, there have been few studies that have separated cytoplasmic and membrane-bound activity. B o m b y x mori fat body contains exclusively particulate GC (Morishima, 1981) whereas silkmoth antennae and M a n d u c a sexta CNS (central nervous system) contain both soluble and particulate activities (Ziegelberger et al., 1990; Morton and Giunta, 1992). The initial

CYCLIC GMP REGULATION AND FUNCTION

3

purification and cDNA cloning of sea urchin and mammalian particulate GC and mammalian cytoplasmic GC revealed the fundamental structural difl'erences between these two types of GC (see Lucas et al., 2000). These are classified as receptor GCs and soluble GCs respectively. Until recently, all subsequently identified GCs could be assigned to one or the other of these classes, but studies reported in the last few years, particularly in invertebrates, have revealed that GCs are a far more diverse family of enzymes than previously thought. 2.1.1

Receptor GCs

2.1.1.1 Sequence analysis. Receptor GCs are integral membrane proteins that were first identified as the proteins associated with the membrane-bound GC activity described above. Seven different isoforms have been cloned in vertebrates. In rats they are named GC-A through GC-G (Garbers and Lowe, 1994; Ffille e! al., 1995; Lowe et al., 1995: Schulz et al.. 1998). Functional studies show that they can be further subdivided into two groups GC-A, GC-B and GC-C that are activated by extracellular ligands, and the retinal GCs, GC-E and GC-F that are activated by intracellular calcium-binding proteins (Lucas et ell., 2000). Sequence comparisons suggest that GC-G belongs to the former group and GC-D the latter (Lucas et al., 2000). BLAST analysis of the Drosophila genome reveals five genetic loci that are predicted to code for receptor GCs and reverse transcriptase-PCR (RT-PCR) with degenerate oligonucleotide primers has identified an additional member of this class in Mamluca, named MsGC-11 (A. Nighorn and D. B. Morton, unpublished data). A schematic diagram of the predicted protein structure of the insect receptor GCs is shown in Fig. 1. The predicted amino acid sequences share the same molecular features: a variable extracellular domain, a single transmembrane domain, an ATP binding/protein kinase-like domain and highly conserved dimerization and catalytic domains. No functional studies have been carried out on the insect receptor GCs and so whether they can be subdivided into the same functional subclasses as the vertebrate receptor GCs remains to be determined. GC-D, GC-E and GC-F are all expressed in sensory neurons, GC-D in olfactory receptor neurons and GC-E and GC-F in photoreceptors (Lucas et ell., 2000). Interestingly, a large family of receptor GCs has been identified in the nematode, Caenorhabdilis elegans, many of which are expressed in olfactory sensory neurons (Yu et al., 1997). The expression patterns of two of the Drosophila receptor GCs have been reported and are not specifically expressed in sensory neurons. Gyc32E is primarily expressed during oogenesis in the ovarioles (Gigliotti et al., 1993) and Gyc76C is expressed in a wide range of tissues, including optic lobe, central brain, thoracic ganglia, digestive tract, oocytes and muscles (Liu et al., 1995; McNeil el al., 1995). Sequence comparison of the insect receptor GCs with the mammalian receptor GCs does not immediately suggest that insect receptor GCs fall into

4

D. B. M O R T O N A N D M. L. H U D S O N

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FIG. 1 Schematic representation of the insect receptor GCs and receptor-like GCs. A Receptor GCs. The insect receptor GCs are shown in comparison with a mammalian receptor GC, rat GC-A. MsGC-II is from Manduca sexta (A. Nighorn and D. B. Morton, unpublished data) and the other receptor GCs are from Drosophila, either previously published sequences (Gyc76C Liu et ell., 1995; McNeil et al., 1995; Gyc32E Gigliotti et al., 1993) or identified from the Drosophila genome sequence using a BLASTP analysis. Each of the five structural domains are shown and are aligned at the predicted transmembrane domain, the position of which was predicted using PSORT II (www.expasy.ch/tools/). The dimerization and catalytic domains were predicted by alignment with GC-A, using CLUSTALW. In the extracellular domains, the vertical lines represent the positions of cysteine residues, and the /x near the transmembrane domain represents the conserved proline-rich juxtamembrane hinge region (see text). Possible ATP binding sites in the kinase-like domain are shown as short, shaded boxes. B Receptor-like GCs. These GCs have catalytic and dimerization domains that are similar to receptor GCs but have no transmembrane or extracellular domains. The only example whose expression has been demonstrated is MsGC-I (Simpson et ell., 1999), but two sequences with a similar predicted structure are present in the Drosophila genome.


5

these two functional groups. Figure 2 shows a phylogenetic dendrogram and reveals that the mammalian "sensory receptor GCs" form a separate cluster that does not include any of the insect receptor GCs. The natriuretic peptide receptors, G C - A and GC-B form a cluster that also contains the Drosophila GC, Gyc32E, and the guanylin/uroguanylin/heat stable enterotoxin receptor, GC-C, forms a separate group with CG4224 and CG3216. The mammalian orphan receptor GC, G C - G , and the other insect receptor GCs tk~rm separate branches from these clusters. Throughout the sequences there are several conserved features and domains, which have been reported to determine specific functional aspects of G C activity. The most extensive structure function analysis has been carried out on GC-A and is used here as the primary comparison. GC-A is a receptor G C that binds and is activated by the peptide hormone, atrial natriuretic peptide (ANP). There is relatively little primary sequence similarity in the extracellular domain between any of the vertebrate and insect receptor GCs, but there are a

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Rat GC-E Rat GC-D

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L CG4224 Rat GC-C CG3216 CG10738 FIG. 2 Phylogenetic dendrogram of the insect and rat receptor GCs. The dendrogram was compiled using the protein sequence parsimony method that infers an unrooted phylogeny (bioweb.pasteur.fl/seqanal/phylogeny/phylip-uk.html).

6

D. B. MORTON AND M. L. HUDSON

number of features that are conserved. The extracellular domains of the mammalian receptor GCs are all of a similar size, varying between 413 and 486 residues. The insect receptor GCs generally have a similar size, 358--476 residues, although CG4224 is predicted to have a much shorter extracellular domain with only 190 residues. The extracellular domain of GC-A contains six cysteine residues that form three intramolecular disulfide bonds (Miyagi and Misono, 2000). The most N-terminal pair forms a disulfide bond between cys-60 and cys-86 (numbering based on the rat GC-A sequence) and is present in all mammalian and insect receptor GCs, with the exception of CG4224, which has a substantially reduced extracellular domain (Fig. 1). The other highly conserved disulfide bond is close to the transmembrane domain (cys423 and cys-432 in GC-A) in a region known as the juxtamembrane hinge (Huo et al., 1999). This region is also notable for being rich in proline residues, in particular pro-419 that is essential for ANP-stimulated GC activity, but not ANP binding (Huo et al., 1999). Replacement of both the cysteine residues in the hinge region with serines also had no effect on ANP binding, but resulted in a constitutively active GC (Huo et al., 1999). These results suggest that this hinge region mediates transmembrane signal transduction for the peptideactivation of GC-A (Huo et al., 1999). Cysteine and proline residues in the equivalent positions are found in all vertebrate receptor GCs, with the exception of GC-C (Huo et al., 1999) and are found in all the insect receptor GCs, with the exception of CG3216 and MsGC-II (Fig. l). All mammalian receptor GCs, except GC-F, contain at least one consensus N-linked glycosylation site (Lucas et al., 2000). All the insect receptor GCs also contain these sites, which vary in number from two in CG3216 to nine in Gyc76C (not shown). Even the reduced extracellular domain of CG4224 contains 6 consensus N-linked glycosylation sites. A number of studies have shown that glycosylation of GC-A, GC-B and GC-C is necessary for ligand binding and activation (see Lucas et al., 2000). The other notable feature of the extracellular domain of GC-A is a binding site for chloride ions in the ANP binding domain, which was revealed when the crystal structure of the GC-A extracellular domain was solved (Van den Akker et al.. 2000). ANP binding is enhanced in the presence of chloride ions (Van den Akker et al., 2000), and it has been suggested that chloride-mediated ANP binding provides a feedback control mechanism for salt regulation (Misono, 2000). Residues that form this chloride-binding site are conserved in CG3216, suggesting that binding and hence activation by its ligand could also be regulated by chloride. Following the transmembrane domain is the kinase-like domain, so called because it contains a consensus ATP-binding site found in all protein kinases (Lucas et al., 2000). However, an essential aspartate residue required for kinase catalytic activity is absent in all receptor GCs (Lucas et al., 2000), including all the insect receptor GCs, and there is no evidence that this domain has kinase activity in any GC. The insect receptor GCs, however, do contain consensus ATP binding sites, although they are not all in the equivalent position to that


7

found in GC-A (Fig. 1). The ATP binding site is an important regulatory domain in mammalian receptor GCs. ATP potentiates the effect of natriuretic peptides on GC-A and GC-B, and alterations in the ATP-binding site eliminate this effect (Duda et al., 1993). In addition, the ATP-binding site in GC-A contains six serine and threonine residues that are phosphorylated in the unstimulated receptor and are essential for peptide-activation of GC-A (Potter and Hunter, 1998). Activation of GC-A by ANP results in dephosphorylation and desensitization of the receptor (Potter and Hunter, 1998). All the insect receptor GCs have one or more serine or threonine residues in close proximity to their consensus ATP-binding site that might act in the same manner. The dimerization and catalytic domains show the highest sequence conservation between mammalian and insect receptor GCs. The dimerization domain, identified in GC-A as a sequence of 42 residues that forms an amphipathic helix (Wilson and Chinkers, 1995), shows 4 7 76% identity with the equivalent region in the insect receptor GCs. Analyzing this region for protein secondary structure reveals that in all the insect receptor GCs this region is also predicted to form an c~ helix (D. B. Morton, unpublished observations). Similarly, the catalytic domain is highly conserved, with the insect receptor GCs sharing 60 69% sequence identity with GC-A. Most importantly, all the residues that have been predicted to form the binding site for the Mg-GTP substrate (Liu et al., 1997) are 100% conserved in all insect receptor GCs. The C-terminal tail of receptor GCs, which immediately follows the catalytic domain, is extremely variable in its length and primary sequence. GC-A and GC-B have no extension beyond the catalytic domain, whereas the other mammalian receptor GCs have an additional 40 60 residues. The insect receptor GCs vary from having no extension in Gyc32E to 443 residues in Gyc76C. Little is known concerning potential functions for this domain, although there is speculation that for GC-C, -E and -F it may be involved in the association of the GC with the cytoskeleton (Lucas et al., 2000). In addition, GC-C undergoes ligand-mediated endocytosis that might be mediated by a YXXZ motif, where Z is L, l, V, M, C or A (Lucas et al., 2000). A similar motif is found in the C-terminal domain of CG4224. Sequence homology between some insect GCs and the natriuretic peptide clearance receptor (NPR-C) indicate a possible novel method of signaling cross talk in these GCs. The NPR-C has an extracellular domain with homology to GC-A, a transmembrane domain and a short cytoplasmic region but no catalytic domain (Lucas et al., 2000). Several studies have shown that this receptor inhibits adenylyl cyclase via activation of Gi (Murthy and Makhlouf, 1999; Pagano and Anand-Srivastava, 2001). The motif that actiw~tes Gi has been identified as containing two basic residues at the amino terminus and a motif that contains BBXXB, BBXB or BXB at the C-terminus, where B represents a basic residue and X is any other residue (Murthy and Makhlouf, 1999: Pagano and Anand-Srivastava, 2001). Interestingly, the Drosophila GCs, Gyc76C, CG10738 and CG9783 also contain these motifs.

8


It will be interesting to determine whether they are capable of activating Gi and hence possibly interacting with an additional signaling cascade. 2.1.1.2 Ligands and activators. There are several peptide hormones that are known to activate mammalian receptor GCs. The natriuretic peptides ANP, BNP and C N P activate GC-A and -B, while G C - C is activated by guanylin, uroguanylin and the bacterial heat stable enterotoxin (Lucas et al., 2000). A BLAST analysis of the Drosophila genome with each of these peptides showed no significant matches (D. B. Morton, unpublished observations). This is not too surprising as there is little similarity in the ligand-binding domains between the mammalian and insect receptor GCs. There are, however, examples of invertebrate peptides that activate membranebound GCs. The crustacean hyperglycemic hormone ( C H H ) activates G C activity in membrane preparations of several crustacean tissues (Goy, 1990; Scholz et al., 1996) and this peptide is part of a large family of related peptides, several of which are found in insects. Figure 3A shows an alignment of the known members of this family found in insects and also includes a sequence identified in Drosophila (CG13586) using a BLAST search with CHH-B. The best characterized of these peptides is the ion transport peptide of Schistocerca gregaria (SgITP) (Meredith et al., 1996) and the almost identical peptide in Locusta migratoria (LmITP). SglTP stimulates salt and water reabsorption in the ileum and acts through an increase in cAMP levels (Chamberlin and Phillips, 1988). A closely related peptide ITP-L is generated by alternative splicing of the m R N A and is not active in the locust ITP ileum bioassay (Meredith et al., 1996) and hence might act through other pathways such as c G M P in this or other tissues. The orthologous Drosophila peptide is part of a predicted protein that is much larger than the locust prohormone (430 compared with 130 residues). This could however be due to inaccurate predictions FIG. 3 Possible ligands and activators of insect receptor GCs. A CLUSTALW alignment of insect peptides that have sequence similarity with the lobster crustacean hypoglycemic hormone B (HaCHH-B). The peptides shown are Bombyx mori CHHlike peptide (BmCHH-L) (Endo et al., 2000), the ion transport and ion transport-like (ITP and ITP-L) of Schistocerca gregaria and Locusta migratoria (Meredith el a/., 1996) and part of a sequence from a predicted protein identified in the Drosophila genome (CG13586). Note that in the predicted Drosophila sequence there is an insertion of 39 residues that is absent in the other sequences. B CLUSTALW alignment of insect GCAP-Iike proteins compared with GCAP-1 from mouse. All of the related proteins are from Drosophila and include frequenin (Frq) (Pongs el al., 1993), neurocalcin (Nca) (Teng et al., 1994) and three genes identified from the Drosophila genome. The predicted N-terminal myristoylation site and the four EF hands arc also indicated. The first EF hand is predicted to be non-functional due to a conserved proline residue at position 4 (see text). The abbreviations for the residues that make each EF hand are as follows, o oxygen in carboxyl side chain (D, N, Q, E, S or T), * any residue, g glycine, j leucine, isoleucine or valine and e glutamate or aspartate.


9

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of the intron/exon structure of the gene, as there are 39 residues in the middle of the predicted mature peptide that are absent in any of the other family members. Several features that are conserved in the peptide family are present in the Drosophila peptide, and include the N-terminal dibasic cleavage site, all six cysteine residues that form three disulfide bonds in CHH and a possible amidation site at the C-terminus of the peptide. Whether this peptide is indeed expressed in Drosophila or whether it activates one of the receptor GCs remains to be determined. The other known activators of receptor GCs are the intracellular, calciumregulated GC-activating proteins (GCAPs) that activate the retinal GCs, GC-E and GC-F (Palczewski et al.o 2000). The GCAPs bind to the retinal GCs and activate them in the presence of low (< I#M) calcium (Dizhoor and Hurley, 1999). BLAST analysis of the Drosophila genome with the GCAP-1 sequence reveals the presence of five closely related genes (Fig. 3B), two of which, frequenin and neurocalcin, have previously been described (Pongs et al., 1993; Teng et al., 1994). The GCAPs and four of the Drosophila GCAP-like proteins have a consensus N-myristoylation site and all contain four EF-hand domains. EF-hand domains are 12 residue calcium-binding loops that contain two helices (Palczewski et al., 2000). Residues at positions 1, 3, 5 and 9 contain an oxygen atom in their side chain (usually from D, N, Q or E but sometimes S or T); position 8 is a leucine, isoleucine or valine and position 12 is glutamate or occasionally aspartate. A glycine residue at position 6 is required for the correct turn between the two helices. The first EF-hand in the GCAPs is nonfunctional, primarily because the proline in position 4 disrupts the formation of the first helix. All five of the Drosophila sequences share these features, as they appear to have a non-functional EF-I and functional EF-2, -3 and -4. The GCAPs are part of a subfamily of myristoylated calcium-binding proteins that contain four EF-hands (Palczewski et al., 2000). This family includes the GCAPs that activate retinal GCs in low-calcium (Palczewski et al., 2000), neurocalcin, which activates GC-E in elevated calcium (Kumar et al., 1999) and recoverin, which inhibits rhodopsin kinase (Palczewski et al., 2000) and does not activate the retinal GCs (Gray-Keller et al., 1993), A more distantly related EF-hand calcium binding protein, SI00B also activates the retinal GCs in elevated calcium (Palczewski e: al., 2000). Hence it is not straightforward to predict which, if any, of the Drosophila proteins will regulate receptor GC activity, and, if they do regulate GC activity, whether it will be in response to elevated or reduced calcium levels. Frequenin was first isolated as a shaker-like mutant that was characterized as having altered synaptic efficacy (Pongs et al., 1993). h~ vitro assays showed that frequenin activated bovine retinal GCs at low calcium concentrations (Pongs et al., 1993). CG5744 is 96% identical to frequenin and would be expected to have similar properties. Drosophila neurocalcin, however, does not show activity in an in vitro assay for retinal GC activation, and has been suggested to function as an inhibitor of G protein-coupled receptor kinases (Faurobert et al., 1996). The observed lack of


11

retinal G C activation could, however, be due to poor species cross reactivity as mammalian neurocalcin will only activate G C - E and has no effect on G C - F ( K u m a r et al., 1999). Hence, it appears that there are candidate G C A P s in Drosophila and presumably other insects, but their respective receptor GCs, and whether they are positively or negatively regulated by increasing calcium levels, remain to be determined. 2.1.2

Soluble GCs

The other major class of GCs are soluble proteins that exist as heterodimers with one c~ and one/4 subunit (Lucas et al., 2000). Activation of these enzymes by extracellular signals requires the generation of an intermediate intracellular messenger, of which the most widespread is believed to be the gas, nitric oxide (NO) (Lucas et al., 2000). Production of NO is achieved by the calciumdependent activation of NO synthase (NOS) (Bredt and Snyder, 1994). NOS and N O / c G M P signaling in insects has recently been reviewed (Davies, 2000; Bicker, 200l). Each soluble G C heterodimer contains a single heme group, which contains a central Fe 2+ or Fe ~+ ion that is coordinated in the plane of the heme by four nitrogen atoms (Lucas et al., 2000). The iron is also coordinated by a histidine residue provided by histidine 105 of the fi subunit (Zhao el al., 1998) that is necessary for NO activation of the GC. Although no spectroscopic studies have been reported on insect soluble GCs, their high degree of sequence conservation and their similar biochemical properties suggest that they bind heine in a similar manner (Liu et al., 1995: Nighorn el al., 1998). 2.1.2.1 Sequence analys& and biochemical properties. Both ~ and fi subunits for soluble GCs from Drosophila, (Liu et al., 1995: Shah and Hyde, 1995) and Mamhwa, (Nighorn et al., 1998) have been cloned and expressed in heteroIogous cells and a fi subunit has been cloned from the mosquito Anopheles ,~ambiae (Caccone et al., 1999). Each of these subunits is similar in size and sequence to their mammalian homologs. The soluble G C subunits can each be divided into two functional domains, a C-terminal catalytic domain that is very similar to the catalytic domain of receptor GCs, and an N-terminal regulatory domain that binds the heine group. Some reports also designate a dimerization domain located between the catalytic and regulatory domain, but this is primarily by analogy with the dimerization domain of the receptor GCs and because the o~and fl subunits do dimerize. There have been no functional studies to show that this region mediates dimerization in soluble GCs, and there is no primary sequence similarity with the dimerization domain of receptor GCs. However, a region of about 40 residues located at the N-terminal side of the catalytic domain shows a high probability of forming a two-stranded coiled coil structure in both the M a m h w a and Drosophila o~ subunits of the soluble GCs (D. B. Morton, unpublished obserw~tions). Coiled coils are common structural

12


motifs that often mediate protein protein interactions and can be predicted using the M U L T I C O I L program (Wolf et al., 1997). Interestingly, neither of the/3 subunits that form heterodimers with these oe subunits shows equivalent coiled coil motifs suggesting that if this region is the dimerization domain, only one subunit needs to form this structure. The crystal structure of the adenylyl cyclase catalytic domain, which has significant sequence similarity to that of GCs, has been solved, and it has been revealed that the active site forms a wreath-like dimeric structure (Zhang et al., 1997). Based on this information, homology modeling has been used to predict the catalytic doraain structure of both soluble GCs and receptor GCs, and has predicted which residues bind the M g - G T P substrate (Liu el al., 1997). The principal difference between the soluble and receptor GCs is that the soluble GCs are heterodimers, whereas receptor GCs are homodimers. One subunit of each dimer binds to the guanosine moiety (the fi subunit for soluble GCs) and the other to the Mg 2+ and triphosphate moieties (Liu et al., 1997). These predictions are shown in Fig. 4, and the conservation of the residues that associate with M g - G T P in the insect subunits is shown in Table 1. Because the soluble GCs are heterodimers, residues in the A strand do not need to be conserved in fi subunits, and conversely, residues in the B strand do not need to be conserved in oe subunits. This is not the case for receptor GCs, where all critical residues are present in both subunits. This arrangement leads to the formation of a single G T P binding site in soluble GCs and two in receptor GCs (Lucas el al., 2000). These active site residues of the insect soluble G C subunits are 100% conserved compared with their mammalian homologs. One deviation in the model compared with the actual sequences is in residue 12 that predicts a serine in the A strand to bind to the }, phosphate, but there is an asparagine at the homologous position of both insect and mammalian ot subunits. The presence of serine in mammalian and insect/4 subunits suggests that the B strand in soluble GCs provides this residue. Similarly, critical residues have been identified in the regulatory region and are also conserved in the five insect soluble G C subunits (Fig. 5). These include histidine 105 in the/4 subunit that forms the axial ligand to the iron of the heme group (Zhao et al., 1998) and two cysteines (at positions 78 and 214 in the rat ill) subunit that are necessary for NO activation (Friebe et al., 1997). One notable difference between the regulatory domains is that although the M a n & l e a regulatory domain contains the same number of residues as mammalian/4 subunits (Nighorn el al., 1998), the Drosophila/4 subunit contains an insertion of 118 amino acids near its N-terminus (Shah and Hyde, 1995). This results in a/4 subunit of 86 kDa, which is larger than the o~ subunit, whereas all other/4 subunits are smaller than the ot subunits. Interestingly, the Anopheles/4 subunit also has an insertion of 91 residues in the same position, although there is no sequence conservation of the inserts between the two flies (Caccone et al., 1999).

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O ......... P

/ o,

, f

..--

;

!"°:;i~,b:::;i[" ...... 3-, ,/" ',..,

"'o\

O

.......... H O / ' ' ' "

9- R 574A

12 S 604A

\ ~o-D 484A E 607A

FIG. 4 Predicted catalytic domain for the soluble GCs. The model for the Mg-GTP binding site is derived from the crystal structure of adenylyl cyclase and is redrawn from Liu el a/. (1997) with permission from The National Academy of Sciences, USA, copyright (1997). The residues that are predicted to be in close association with Mg-GTP are numbered, clockwise l¥om 1 to 17, starting from the arginine at position 53l. The position in either the A strand or B strand is indicated and the amino acid number is given for the Man&~ca soluble GC, either MsGC-oH for the A strand or MsGC-fll for the B strand.

14

TABLE 1


Residues in the G T P binding d o m a i n of the insect soluble G C s

A Strand position

6

7

9

10

11

12

14

Model prediction

T

D

R

D

E

S

I

-

N N -

DGC-c~ 1 MsGC-cel DGC-fl I MsGC-fil A g G C-fi 1 MsGC-fi3 CG4154 CG14885 C G 14886

. O A G

.

. -

. T T T

O O G

M M

Y Y

-

E E

V V V V V

-

B Strand position

1

2

3

4

5

8

13

15

16

17

Model prediction

R

C

F

G

V

N

R

F

M

E

D G C - / ~ I

-

MsGC-~I AgGC-~I DGC-ce I MsGC-oel MsGC-fi3 GC4154 GC14885 G C 14886

K

-

. -

.

. -

-

T T

K K -

C C

. -

The positions of each of the residues and their predicted interactions with GTP are shown in Fig. 4. A dash indicates that the residues in the insect GCs are identical to the mammalian receptor GC, GC-A, which was used in the original rnodeling study (Liu et al., 1997).

T h e i n s e c t s o l u b l e G C s u b u n i t s also s h o w s i m i l a r b i o c h e m i c a l p r o p e r t i e s t o t h e i r m a m m a l i a n h o m o l o g s w h e n e x p r e s s e d in h e t e r o l o g o u s cells. T h e s e e x p e r i m e n t s h a v e b e e n c a r r i e d o u t w i t h b o t h t h e Drosophila ( S h a h a n d H y d e , 1995) a n d M a n d u c a ( N i g h o r n et al., 1998) s u b u n i t s . B o t h , like t h e m a m m a l i a n e n z y m e s , a r e o b l i g a t e h e t e r o d i m e r s , i.e. n o a c t i v i t y is s e e n if e i t h e r s u b u n i t is e x p r e s s e d o n its o w n , b u t a c t i v i t y c a n b e m e a s u r e d if b o t h s u b u n i t s a r e c o t r a n s f e c t e d i n t o cells. T h i s a c t i v i t y w a s s t i m u l a t e d in t h e p r e s e n c e o f N O d o n o r s . I n a d d i t i o n , t h e M a n d u c a h e t e r o d i m e r h a d a h i g h e r level o f b a s a l a c t i v i t y a n d l o w e r N O - s t i m u l a t e d a c t i v i t y w h e n M n - G T P w a s u s e d as a s u b strate compared with Mg-GTP, and NO-activation was blocked by the soluble GC inhibitor ODQ (lH-[1,2,4]oxadiazolo[4,3-a]quinoxalin-l-one) (Nighorn et al., 1999). B o t h o f t h e s e c h a r a c t e r i s t i c s h a v e also b e e n d e s c r i b e d f o r m a m m a l i a n s o l u b l e G C s ( L u c a s et al., 2000).

CYCLIC GMP REGULATION AND FUNCTION Cys-78

RatGC ~i 76 MusGC ~i 76 HumGC-~I BovGC-~I

medGC MsGC-~I Dgc-~

AgGC ~i MsGC-~3 CG4154 CG14886 CG14885

76 76 76 76 76 75 75 75 75 57

15 Cys- 214

His-105 203 203 203 203 203 201 323 294 198 198 215 197

KHLPi~ASVLFEI~ KHLPI~AHVLFEIF~ KMPS~DLNVFLELF~P KMPT~KLDVFLDLFP

FIG. 5 Alignment of portions of the regulatory domain of the soluble GC [4 and filike subunits from vertebrates and insects. The residues highlighted in black are the highly conserved cysteines (78 and 214) and histidine 105. Gray highlighting shows residues that are conserved (identical and similar) between vertebrates and at least one insect GC. Sequences shown for vertebrate GCs are for the fll subunits of rat, mouse (mus), human (hum), cow (boy) and medaka fish (reed). Insect sequences are flom Manduca (MsGC-fil and MsGC-fl3), Anopheles (AgGC-fil) and Drosophila (Dgc-fl, CG45154, CG14886 and CG14885).

A BLAST analysis of the Drosophila genome also identified three additional GCs that had significant sequence similarity with /-,{ subunits of soluble GCs. These are discussed further, together with a unique Manduca fi subunit, named MsGC-/-]3, that appears to be a member of a separate class of GC.

2.1.3

Atypical GCs

As described above, GCs have classically been divided into two classes receptor and soluble GCs. Recent studies, however, have identified three additional types of GC, two of which have been identified in Man&tea. One, named MsGC-I, is related to receptor GCs, but is not an integral membrane protein and lacks the extracellular and transmembrane domains characteristic of receptor GCs (Simpson el al., 1999). The other, named MsGC-fl3, is related to soluble GCs but can function as a homodimer and is NO-insensitive (Nighorn et al., 1999). The third new class of GC is structurally very different. It is a bifunctional protein that has an N-terminal ATPase domain and a Cterminal domain that is structurally similar to adenylyl cyclases containing 12 transmembrane domains (Linder el al., 1999). The C-terminal adenylyl cyclaselike domain contains catalytic domains similar to GCs, and recombinant proteins have GC activity and no adenylyl cyclase activity (Carucci et al., 2000; Linder et al., 2000). GCs with this structure have been found in the protozoa Paramecium, Tetrahynwna and Plasmodium. Recently, a GC was also cloned from Dictoslelium that also had a predicted topology similar to adenylyl cyclases (Roelofs et ell., 2001). There is no evidence that similar GCs are

16


f o u n d in insects or other multicellular organisms a n d these will not be discussed further. A s u m m a r y of the different classes of G C a n d their phylogenetic d i s t r i b u t i o n is shown in Figure 6.

Ligandbinding

Heme-

Trans- ~ . membrane

~ase i -like

Regulatory?

||

Dimerization Catalytic

~

binding

II A

B

II

C ATPaseDomain

GC Domain

D

j u U v ouov E

Catalytic Domain

FIG. 6 Schematic diagram of the different classes of GCs identified in insects and other organisms. A Receptor GCs are homodimeric integral membrane proteins with ligand-binding, transmembrane, kinase-like, dimerization and catalytic domains. They have been identified in insects, mammals, fish, birds, sea urchins, and C. elegans. B NO-sensitive soluble GCs are heterodimeric cytoplasmic enzymes with heme-binding and catalytic domains. They have been identified in insects, mammals and fish and are predicted to be absent in C. elegans (Morton et al., 1999). C Receptor-like GCs are similar to receptor GCs, but lack the ligand-binding and transmembrane domains. The only definitive example is in Manduca, but genomic sequences exist that predict their presence in Drosophila, C. elegans and the acidian Ciona hltesthmlis. D NO-insensitive soluble GCs are active as homodimers and have a similar domain structure to NO-sensitive soluble GCs, but are not activated by NO. Manduca is the only organism where conclusive evidence for their presence has been demonstrated, but they are predicted to also be present in Drosophila and C. elegans. E Multiple transmembrane domain GCs have been found in the protozoa and Dictostelium. The protozoan GCs have an ATPase domain and a GC domain with 12 transmembrane domains, whereas the Dictostelium GC only has the GC domain.


17

M s G C - I and related GCs. MsGC-I was isolated using RT-PCR and degenerate oligonucleotides designed to hybridize to regions of the catalytic domain that are conserved between soluble and receptor GCs (Nighorn et al., 1998). Sequencing the PCR product suggested that MsGC-I was a receptor GCs; however, isolation and sequencing full-length cDNAs showed that it did not contain a signal sequence, extracellular or transmembrane domains (see Fig. 6). Northern blots showed that the eDNA was similar in size to the 2.5 kb transcript, suggesting that an incomplete cDNA had not been cloned. In addition, antisera made to a fusion protein recognized a 55 kDa protein on Western blots of nervous tissue the size of protein predicted from the sequence (Simpson et ~ll., 1999). The catalytic domain is 77% identical to the catalytic domain of GC-B, a mammalian receptor GC, compared with 33% identity with soluble GCs, and contains all the residues necessary for Mg-GTP binding. In addition, MsGC-I contains a region that is 76°/,, identical to the dimerization domain of GC-B (Simpson et al., 1999). A schematic diagram of the domains of MsGC-I is shown in Fig. lB. MsGC-I contains a C-terminal extension that shows no homology to any GC or any other protein in the databases, and a short Nterminal extension that shows a low level of similarity to receptor GC kinaselike domains but does not contain a consensus ATP binding site. There is no similarity to the regulatory domain of soluble GCs. MsGC-1 belongs to a new class of GC that is related to receptor GCs, but is not an integral membrane protein and cannot be activated directly by an extracellular ligand. There is a report of a mammalian GC cloned from rat kidney, ksGC, that also has the same structure as MsGC-I (Kojima et al., 1995). No GC activity was measured t¥om recombinant ksGC, and there was no evidence that a protein of the predicted size was indeed expressed in vivo. A subsequent report has suggested that ksGC was a cloning artifact and was a partial clone of the receptor GC, GC-G (Schulz et al., 1998). When MsGC-I was transiently expressed in COS-7 cells it showed a high level of basal GC activity, was not stimulated by NO, and was located in the soluble traction of COS-7 cell homogenates (Simpson et al., 1999). Further experiments with recombinant MsGC-I showed that it formed homodimers when expressed in COS-7 cells (Simpson et al., 1999). Although it was located in the soluble fraction of COS-7 cells, it was present in the particulate fraction of Manchu'a nervous tissue homogenates, suggesting that it was bound to another protein in vivo (Simpson e t a / . , 1999). hi situ hybridization and immunocytochemistry showed that MsGC-I was expressed in a small population of neurons in the posterior of each segmental ganglion of pre-pupal animals (Simpson et al., 1999). Analysis of the distribution of MsGC-I has also been extended to the brain and antennae of adult M a m h u ' a . MsGC-I showed extensive expression throughout the brain, especially in sensory neuropil of the olfactory and visual systems and the higher order neuropil of the mushroom bodies and central complex (Nighorn et al., 2001 ). Expression of MsGC-I was also seen in the cell

2.1.3.1

18


bodies and dendrites of the olfactory sensory neurons in the antennae (Nighorn et al., 2001). Several studies have noted the role of N O / c G M P signaling in olfaction and olfactory processing in insects (see section 3.2.1). The distribution of MsGC-I in the olfactory system raises the possibility that there is a parallel pathway of NO-independent signaling, although co-localization of both NO-sensitive GCs and MsGC-I in the same cells has yet to be demonstrated. A major question to address is the mechanism of MsGC-I regulation. It is not activated by NO and it cannot be directly activated by extracellular ligands. When MsGC-I was expressed in COS-7 cells and the accumulation of c G M P in intact cells measured, the level of c G M P present was higher compared with cells that were transfected with the NO-sensitive soluble GC and stimulated with NO (Simpson et al., 1999). It seems unlikely that MsGC-I exhibits this high level of basal activity in ~,ivo, unless it is co-expressed with a constitutively active PDE. As MsGC-I has a different subcellular distribution in COS-7 cells (cytoplasmic) compared with its distribution itl ri~,o (particulate fraction), it is tempting to speculate that iu vivo MsGC-I interacts with a membrane associated protein (possibly a receptor) and its activity is inhibited in this state. Activation could then be achieved by dissociation from this protein, yielding an active GC. Another possible mechanism of activation has been revealed recently by examining the expression of MsGC-I in peripheral neurons. The epidermis of M a n d u c a is innervated by a population of dendritic arborization neurons that are thought to be mechanosensory (Grueber and Truman, 1999). Most of these neurons responded to NO donors with an increase in c G M P (Grueber and Truman, 1999), and hence probably contain MsGC-od and MsGC-/?I. One specific neuron, ddaB, however, responded very weakly to NO but showed a large cGMP increase ill the presence of EGTA, which was assumed to reduce the levels of cytoplasmic calcium in the cell (Grueber and Truman, 1999). This neuron specifically stained with the antisera to MsGC-I, suggesting that a reduction in calcium might be involved in its activation in ~'iro (Grueber et al., 2001). This might involve a similar mechanism of activation described for the receptor GCs mediated by the GCAPs in mammalian retina (Dizhoor and Hurley, 1999, see section 2.1.1.2). In support of this model we showed that antisera that recognize the Drosophila GCAP ortholog frequenin, also stained ddaB (Grueber et al., 2001). MsGC-I is the only reported example of a GC with its particular structural characteristics whose in vivo expression has been demonstrated. Genetic loci that are predicted to code ['or proteins with similar structures to MsGC-I are present in C. elegans, Drosol)tfila and the Acidian Ciona intes'tinalis (D. B. Morton, unpublished observations). Although these sequences suggest that MsGC-I is not unique, experimental evidence of their expression is needed. Schematic diagrams of the two predicted Drosophihl proteins, CG5719 and CG9783 are shown in Fig, lB. The catalytic domains, like that of MsGC-I,


19

are more similar to those of receptor GCs than soluble GCs, and all the residues involved in M g - G T P binding are again 100% conserved. Both CG5719 and CG9783 have regions with similarity to the dimerization domains of receptor GCs and both are predicted to form ~ helices. The two Drosophih~ sequences have a region N-terminal to the dimerization domain that is larger than the one found in M s G C - I and is similar in size to the kinase-like domains of receptor GCs. A search for protein motifs in this region of CG9783 identifies it as being similar to protein kinases, and a BLAST analysis shows that it has about 45% identity to the kinase-like domain of several mammalian receptor GCs. It contains a consensus ATP binding site that has two serine/threonine residues nearby that could be phosphorylated in a similar fashion to GC-A (Potter and Hunter, 1998). This region also lacks the critical aspartate residue present in all protein kinases but lacking in the kinase-like domain of all receptor GCs. By contrast, the N-terminal portion of CG5719 shows no similarity to the kinase-like domains of receptor GCs and shows no homology to any other protein. Additional studies are clearly required on the members of this novel class of G C to determine whether their predicted structures are correct, how they are activated and what role they play in regulating c G M P levels in specific cells.

2.1.3.2 MsGC-fi3. During the course of the study that identified the Mandm'a GCs, MsGC-c~I, MsGC-fll and MsGC-I, we isolated an additional G C that was also closely related to mammalian fi subunits. Two mammalian G C fi subunits have been identified, fil and fi2. The new Manduca fi subunit had a number of specific differences that led us to believe that it was not a homolog of either the fil or fi2 subunits, and we named it MsGC-fi3 (Nighorn et al., 1999). The principal novel structural feature that MsGC-fi3 contains is a C-terminal extension of 315 residues that has no similarity to any other protein in the databases (see Fig. 6). There are no identifiable domains in this Cterminal sequence, but there is a predicted C-terminal isoprenylation site and several possible protein phosphorylation sites throughout its length (Nighorn el al., 1999). Although the regulatory domain of MsGC-fi3 is similar to other/31 subunits (both insect and vertebrate), there are a number of specific differences. The most notable are the lack of two cysteines (78 and 214) that are 100% conserved in all other fl subunits and are necessary for NO activation (Friebe et al., 1997) (Fig. 5). This suggested that MsGC-fi3 would form a G C that is insensitive to NO activation. It does, however, contain a histidine at the equivalent position of histidine 105, believed to be the axial ligand for the heme group in soluble GCs (Zhao el al., 1998). The catalytic domain of MsGC-fi3 also revealed some interesting features. All of the residues that are predicted to associate with the M g - G T P substrate are conserved in MsGC-fi3, here it is more similar to the homodimeric receptor

20


GCs than the heterodimeric soluble GCs (Table 1). The A strand residues 6-T, 9-R and 10-D that are replaced by non-conservative substitutions in all other fll subunits are conserved in MsGC-fi3 and the B strand 8-N and 16-M that are replaced in all other c~ subunits are also conserved in MsGC-fiY This is particularly noteworthy for 6-T and 8-N that are involved in the condensation reaction of the 3' hydroxyl of the ribose to the oe phosphate group and for 9R and 10-D that are involved in stabilization of the metal-triphosphate moiety (Fig, 4). Thus MsGC-fl3 provides all the residues needed for binding Mg-GTP without the need of a second subunit, suggesting that it might be able to form active homodimers. Transient expression of MsGC-/~3 in COS-7 cells showed that both of the predictions based on sequence analysis were valid. Although all the NOsensitive soluble GCs that have been examined are obligate heterodimers, MsGC-fi3 showed significant basal activity in the absence of additional subunits (Nighorn et al., 1999) and subsequent gel filtration data strongly supported the prediction that MsGC-fi3 formed homodimers (D, B. Morton, unpublished data). When MsGC-fl3 was co-expressed with either of the other M a n d u c a soluble G C s , MsGC-oel or MsGC-fll, the total level of enzyme activity was slightly reduced, suggesting that heterodimers were formed and that they had either reduced or negligible activity compared with homodimeric MsGC-fl3 (Nighorn et al., 1999). The activity of MsGC-fi3 was also insensitive to activation by NO (Nighorn et al., 1999). Like other GCs, MsGC-fl3 showed higher levels of activity when manganese was present, as compared with magnesium, but in neither case was the activity stimulated by NO. In addition, co-expression with either MsGC-otl or MsGC-fil did not yield a NO-sensitive GC (Nighorn et al., 1999). These properties have not been reported for any other native GC, but when the rat/41 subunit was mutated to remove cysteines 78 and 214, and subsequently co-expressed with wild type rat otl, a NO-insensitive GC was produced (Friebe et al., 1997). In this case, however, NO-sensitivity could be restored by incubating the mutant [31/oel G C with excess heine (Friebe et al., 1997). By contrast, when heine reconstitution experiments were carried out with either MsGC-/43 expressed alone or co-expressed with MsGC~1, the activity was still unaffected by NO donors (Nighorn e t a / . , 1999), It is not clear whether MsGC-fi3 is unable to bind heme, or whether it does so in a configuration that renders it insensitive to NO. The soluble GC inhibitor, 1H[1,2,4]oxadiazolo[4,3-a]quinoxalin-l-one (ODQ), acts on NO-sensitive soluble GCs by oxidizing the ferrous heme to ferric heme (Zhao et al., 2000) and we found that MsGC-fl3 was insensitive to ODQ at concentrations up to 100 # M (Nighorn et al., 1999). This suggests that MsGC-fl3 does not require ferrous heme for activity, but spectroscopic studies will be needed to unequivocally determine the presence and nature of the heme group. The unusual properties of this novel GC raise several important questions. These include understanding the physiological functions that it mediates, its mechanism of activation and whether it forms homodimers or heterodimers in


21

viro. A significant contribution to these issues came from the finding that MsGC-fi3 is expressed in cells that respond to the neuropeptide, eclosion hormone (EH). EH is a central component of the endocrine cascade that triggers ecdysis behavior (shedding of the old cuticle) at the end of the molt (see Chapman, 1998). A series of studies (discussed in more detail in section 3.2.3) have shown that EH acts through an increase in c G M P and probably activates a novel NO-insensitive soluble GC (Morton and Truman, 1985: Morton and Giunta, 1992; Morton 1996; Kingan el al., 1997). Three specific EH target cell populations have been identified: a population of about 50 neurons in the ventral nerve cord (VNC) that contain crustacean cardioactive peptide (CCAP) (Ewer et al., 1994), the Inka cell in the peripheral epitracheal glands (Zitnan el al., 1996) and a population of non-neuronal cells in the transverse nerve of the abdominal ganglia (STNR cells) (Hesterlee and Morton, 2000). RT-PCR experiments showed that MsGC-fi3 was expressed in the transverse nerve and the epitracheal glands (Nighorn et al., 1999) and immunocytochemistry showed expression of MsGC-fi3 in the STNR and Inka cells (Morton, 2000). These studies provide circumstantial evidence for the activation of MsGC-fi3 by EH, but as the sequence of MsGC-fi3 predicts that it is cytoplasmic and as EH is an extracellular hormone, this activation must involve intermediate signals. Possible pathways for this activation are discussed in section 3.2.3. MsGC-fi3 is the only GC with the sequence characteristics described above that has been demonstrated to be a NO-insensitive GC. There are, however, several other GCs with these same features present in a variety of organisms. The nematode, Caenorhabditis elegasls has seven genes that code for soluble GCs and none of them have a cysteine in the equivalent positions as cys78 and cys214 (Morton el al., 1999). The lack of an identifiable NOS gene in the genome of C. eh,,gans (Bargmann, 1998) and the finding that GC activity in ('. el

m ~o r -

H

m

~,~ SS

C~ ~Z

m

a

H

a

H

m ~

m ~0 co en ~n m

~ m Lrt ~0 r--

O4

H m

0 m

,~ ca Z

~ o ~ .~NNNN~

rn

,

¢aj

,~ . ~

,~

26


Several of the PDE families contain G A F regulatory domains (named after the cGMP-binding PDEs, the Anabaena adenylyl cyclase and the fhlA gene) that form allosteric binding sites for c G M P in addition to the catalytic site (Soderling and Beavo, 2000). These domains have been studied in most detail in PDE5, where they appear to be involved in a negative feedback loop for regulating cGMP levels. PDE5 contains two c G M P binding sites that are required for its phosphorylation by both cAMP-dependent protein kinases (PKA) and cGMP-dependent protein kinases (PKG) (Turko et al., 1998b), and this phosphorylation increases the catalytic activity of PDE5 (Corbin et al., 2000). As the levels of cGMP increase in a cell, c G M P first binds to the catalytic site, which increases the affinity of the G A F binding sites. This allosteric binding exposes the phosphorylation site of PDE5 to PKG, which is also activated by increased c G M P levels. PDE5 is then activated by PKG phosphorylation, bringing the c G M P back to resting levels (Corbin et al., 2000). PROSITE analysis shows that both CG10231 and CG8279 contain two predicted G A F domains. A number of the residues required for c G M P binding have been identified (Turko et al., 1996) and form a NKX,,D motif that is conserved in both G A F domains of CG10231 and CG8279 (Fig. 7B). In addition, a serine in a similar position to the set92 that is phosphorylated in PDE5 is also present in both the Drosophila sequences. This suggests that both CG10231 and CG8279 in addition to regulating the levels of c G M P may be downstream effectors that bind c G M P and might be stimulated by PKG phosphorylation.

3 3.1

Cyclic GMP function MOLECULARTARGETS

There are three primary protein families that act as c G M P receptors within cells. In addition to the cGMP regulated PDEs (see section 2.2), these include cGMP-dependent protein kinases (PKG) and cGMP-gated ion channels. All three families are characterized by containing cyclic nucleotide binding sites. A search of the Protein Family (Pfam) databases for cyclic nucleotide (cNMP) binding sites in insects revealed 17 separate protein sequences (D. B. Morton, unpublished data). All but one (a voltage and cyclic nucleotide-gated ion channel in Heliothis virescens) were from Drosophila. Five of the sequences were protein kinases (two for the regulatory subunits of PKA and three PKGs), nine were ion channels and three additional proteins were identified. Two of these additional proteins were similar to cAMPdependent guanine nucleotide exchange factors (Kawasaki et al., 1998) and the third was the protein product of the swiss cheese (sws) gene (Kretzschmar et al., 1997). Mutants of the sws gene were isolated in screens for structural brain defects and all five known alleles show age dependent


27

neurodegeneration (Kretzschmar et al., 1997). The protein product of sws has three N-terminal cyclic nucleotide binding domains that are most similar to those domains in PKA regulatory subunits (Kretzschmar et al., 1997). However, a multiple sequence alignment of all cNMP-binding sites in insects showed that they were also closely related to the c N M P binding sites of PKGs (D. B. Morton, unpublished observations). The relative affinities of sws for cAMP and c G M P are not known. The importance of the cNMP binding site for normal sws function was demonstrated by the finding that one of the sws mutant alleles contained a point mutation that substituted a glycine that is present in all 17 of the insect cNMP-binding sites. Thus sws probably represents an additional molecular target for cNMPs and might be a target for cGMP. 3.1.1

Proteh7 kinases and substrates

Both mammals and insects have two distinct families of protein kinases that are activated by cyclic nucleotides. The cAMP-dependent protein kinases are tetramers consisting of two catalytic subunits and two cAMP-binding regulatory subunits, whereas the PKGs are homodimers, with each monomer containing both catalytic and cGMP-binding regulatory domains (Siegel el al., 1994). Protein kinase activities that are preferentially regulated by c G M P have been described in a variety of insect tissues, including ttyalophora cecropia body wall (Kuo et al., 1971), fat body from H. cecropia, Antheraeapolyphemus, Mamtuca and the cockroach Blaberus discoMalis (Kuo et al., 1971), B o m b v x pupae (lnoue et al., 1976) and eggs (Takahashi, 1985), Manduca CNS (Morton and Truman, 1986) and whole animal extracts from Drosophila (Kuo et al., 1971) and Ceratitis capitata (Haro et al., 1983). Two genes for P K G have been cloned from Drosophila (Kalderon and Rubin, 1989) and a third gene that appears to code for a PKG has also been identified fi'om the Drosophila genome (Morrison et al., 2000). A BLAST analysis shows that the two cloned PKG genes, Pkg21D (also known as DG1) a n d j b r a g i n g (fbr, also known as DG2), are most similar to mammalian PKG type I and the third, CG4839, is most similar to mammalian P K G type II. All three Drosophila PKGs have a similar domain structure to mammalian PKGs (Fig. 8). The N termini of the proteins are the most variable, but all three appear to have a dimerization domain that contains a leucine/isoleucine zipper motif (Foster et al., 1996). The central region of the predicted protein contains two cyclic nucleotide-binding domains and there is a C-terminal protein kinase catalytic domain. The./br P K G has four different splice variants (Kalderon and Rubin, 1989) that utilize three different initiation codons. The splice variants differ primarily in the N-terminus, although they all contain leucine/isoleucine zippers and hence are predicted to form dimers. Two of the splice variants have a shortened first cyclic nucleotide-binding domain, and hence might only contain one functional cGMP-binding site.

28

D. B. MORTON AND M. L. HUDSON cNMP LZ

1

2

PK

for-P1 for-P2

for-P3 for-p4

[] ........

~

~

Pkg21 D CG4839

[]

~

~

l

n

cGK-I cGK-II

~

l

n

FIG. 8 Schematic representation of the insect PKGs. The domains shown are the leucine/isoleucine zipper (LZ), the two cNMP binding sites and the kinase catalytic domain (PK). The insect PKGs shown are the four splice variants ofjor, Pkg21D and the predicted PKG CG4839. Note that in./br-P3 and/br-P4 exons within the first cNMP-binding domain are absent. Also shown are the two mammalian PKGs, cGKI and cGKII. The biochemical properties of Pkg21D have been investigated and were similar to the properties of mammalian PKGs (Foster et al., 1996). Recombinant Pkg21D formed homodimers, was preferentially activated by c G M P and was autophosphorylated in the presence of c G M P (Foster et al., 1996). Activation of Pkg21D with c G M P showed positive cooperativity, probably reflecting the presence of two cGMP-binding sites (Foster et al., 1996). Although no biochemical investigations have been carried out on t h e / o r gene product, there is experimental evidence, rather than only sequence homology, that suggests it does code for a PKG. The jbr gene was first identified as a naturally occurring polymorphism for food-search behavior (Sokolowski, 1980) and subsequently shown to code for a predicted PKG (Osborne et al., 1997) (see also section 3.2.3). Two naturally occurring alleles o f j o r showed different levels of expression o f J o r mRNA and protein product (Osborne et al., 1997). Correlated with the levels of tbr expression, different levels of cGMPdependent protein kinase activity were measured in animals expressing the different alleles (Osborne et al., 1997), providing circumstantial evidence that Jbr does code for a functional PKG. Relatively few studies have directly investigated specific substrates for PKGs in insects. This is partly because there is considerable overlap between substrate specificities for PKG and PKA. In most tissues, the levels of PKA are much


29

higher than PKG and hence it is difficult to identify specific substrates For PKG. For example, in silkworm eggs, vitellin was shown to be phosphorylated by highly purified preparations of both PKA and PKG, and the K,, for both enzymes was similar (Takahashi, 1985). During early development, the levels of PKG was higher than PKA (Takahashi, 1985) and hence vitellin could have been the endogenous substrate, although no evidence was presented that vitellin was phosphorylated in vivo. In M a m t , ca ventral nerve cords (VNCs) two proteins have been described that are phosphorylated specifically in response to c G M P and appear to be phosphorylated by the action of eclosion hormone (EH) in vivo (Morton and Truman, 1986). These proteins, named the EGPs (EH and cGMP regulated Phosphoproteins) both have a mass of 54 kDa but differ in isoelectric point (about 5.6 and 6.6) (Morton and Truman, 1986). In VNC homogenates, cAMP was more effective than c G M P at stimulating their phosphorylation, but in intact nervous tissue the reverse was true, with c G M P being more effective at stimulating their phosphorylation than cAMP (Morton and Truman, 1988a). This difference is probably because both PKA and PKG can phosphorylate the EGPs, and as levels of PKA are about 10 fold higher than PKG in Manduca VNCs (Morton and Truman, 1986), PKA will preferentially phosphorylate the EGPs in homogenates. This is supported by the finding that partially purified preparations of the EGPs can be phosphorylated by mammalian PKA (D. B. Morton, unpublished data). Presumably, in the specific cells that express the EGPs there are either comparable levels of PKA and PKG, or there is more PKG than PKA, and hence in the intact tissue c G M P is more effective. The EGPs were also phosphorylated by exposure of intact VNCs to EH (Morton and Truman, 1988a). EH acts on the VNC to stimulate an increase in cGMP but has no effect on the levels of cAMP (Morton and Truman, 1985), providing further evidence that the EGPs are specific substrates for PKG. The identity. function and cellular location of the EGPs remain to be determined. Studies on mammalian cGMP-stimulated protein phosphorylation have identified several specific substrates for PKGs (see Smolenski er al., 1998). Several of these appear to be mammalian specific as BLAST analysis of the Drosophihl genome fails to identify orthologs (D. B. Morton, unpublished observations). These include the protein phosphatase inhibitors G-substrate and DARPP-32 and the inositol trisphosphate receptor (IP~R) associated PKG substrate (IRAG). Orthologs of other protcins, shown in mammalian systems to be substrates for PKG, are present in Drosol)hihl and hence are possible substrates for one or more of the Drosophila PKGs. These include the IP3R, L-type calcium channels, calcium sensitive potassium channel, phospholipase C and the cystic fibrosis transmembrane conductance regulator. Another family of proteins that includes substrates for PKG in mammals is of particular interest because of the relationship between c G M P and axonal outgrowth (see section 3.2.2). Many studies have shown that c G M P is a major regulator of vascular smooth muscle relaxation (see Lincoln el a/., 1994) and

30


the vasodilator-stimulated phosphoprotein (VASP) is a substrate for PKG in smooth muscle, vascular endothelium and platelets (Smolenski et al., 1998). VASP is a member of a family of related proteins that includes the product of the Drosophila enabh, d (enb) gene, the mammalian ortholog of enb, mena, and an additional related protein, Ena/VASP-like protein (EVL) (Gertler et al., 1996). The Drosophila enb gene was first identified as a suppressor of the tyrosine kinase, Abels, on (Gertler et al., 1995). Mutations in the enb gene include defects in neuronal dendritic and axonal outgrowth and branching (Gao et al., 1999; Wills el al., 1999) and interacts with robo, a Drosophila axon guidance receptor (Bashaw et al., 2000). BLAST analysis of the Drosophila genome has identified another member of this family, CG10155 (D. B. Morton, unpublished observations). There is accumulating evidence that members of the Ena/VASP family can act as adapter proteins linking extracellular signaling pathways to actin polymerization (Prehoda el al., 1999) and that phosphorylation of VASP regulates its interactions with actin (Harbeck et al., 2000). The functional conservation in the Ena/VASP family has been demonstrated by the rescue of lethal enb mutations with human VASP (Ahern-Djamali et al., 1998), although it is not known whether VASP will rescue dendritic and axon guidance defects. The cGMP-dependent phosphorylation sites of VASP are conserved in mena and EVL (Gertler et al., 1996). Although there is no PKG phosphorylation site (two basic residues 2 and -3 of a serine or threonine) at the equivalent position in the ent~ and CG10155 proteins, there is at least one potential PKG site elsewhere in both sequences. There is accumulating evidence that cGMP plays a critical regulatory role in neuronal outgrowth and pathfinding (section 3.2.2) and hence it is intriguing that the protein product of enb (a gene that affects dendritic and axonal outgrowth and branching) and CG 10155 could be substrates for PKG. 3.1.2 ()'clio nucleotMe-gated channels The Drosophila genome contains predicted genes for three classes of ion channels that have intracellular cNMP-binding domains (Littleton and Ganetski, 2000). These classes are the cNMP-gated ion channels that are orthologs of the mammalian retinal cGMP-gated ion channels (Zagotta and Siegelbaum, 1996), the Ih channels that are activated by hyperpolarization and contain a cNMP-binding site (Ludwig et al., 1998) and the EAG class of voltage activated potassium channels that also contain a cNMP-binding site (Briiggeman et al., 1993). There are four genes in the Drosophilci genome that appear to code for cyclic nucleotide-gated ion channels (CNGs) (Littleton and Ganetski, 2000). Two have been cloned and partially characterized, cyclic nuch, otide-gated ion channel protein (cng) (Baumann el al., 1994) and cng-like (cngl) (Miyazu et al., 2000), while two additional genes, CG3536 and CG17922, have been identified from sequencing the Drosophila genome. The mammalian CNG


31

channels are composed of two homologous (oe and /3) subunits that form tetramers in vivo. although the stoichiometry is not known (Zagotta and Siegelbaum, 1996). Mammalian c~ subunits form functional homomeric channels in heterologous expression systems and although the /3 subunits do not form functional channels on their own, heteromeric channels formed from o~ and /3 subunits have distinct kinetic properties compared to homomeric oe channels (Finn el al., 1998). The cng channel has been expressed in heterologous cells and functions as a homomeric cGMP-gated non-specific cation channel that is 50-fold more sensitive to c G M P than to cAMP (Baumann el al., 1994). The cngl channel, by contrast, does not l~rm a functional homomeric channel in heterologous cells, suggesting that it is more similar to a /3 subunit (Miyazu et al., 2000). Varnum et al. (1995) have identified an aspartate residue near the C-terminal end of the cNMP-binding site that determines the selectivity towards c G M P in the bovine rod CNG. A C L U S T A L W sequence alignment of the predicted cNMP binding sites in the four Drosophila CNGs showed that the c~Tgchannel has an aspartate at the equivalent position, but cngl, CG3536 and CG17922 have valine, tyrosine and asparagine residues in place of the aspartate (D. B. Morton unpublished data). Varnum el al. (1995) found that replacing the negatively charged aspartate with an uncharged polar residue made the channel non-selective towards c G M P and cAMP, whereas replacement with a non-polar residue made the channel selective towards cAMP. Thus, CG3536 and CG17922 would be predicted to have little selectivity towards c G M P or cAMP, whereas cngl would be expected to form a channel specific to cAMP. Finn et al. (1998) report cloning a Drosophila CNG /3 subunit that does not form active homomeric channels. This subunit is likely to be the same as the CG3536 channel, as their sequences are 97.5% identical (D. Krautwurst. personal communication). Co-expression of the /3 subunit with the rat oe subunit of the olfactory CNG (rOCNGc0 yielded a channel that had a larger cGMP-induced current than cAMP-induced currenL whereas homomeric rOCNGoe channels yielded similar cGMP- and cAMPinduced currents (Finn eta/., 1998). This suggests that CG3536 might confer c G M P selectivity to the subunit with which it forms channels in fifo. The cellular expression patterns of cn~ and oHg/ suggest that these two subunits do not form heterodimers in s,ivo. The cHg channel is expressed in the antennae and eyes. although a snore detailed analysis of the expression pattern in specific cells is not known (Baulnann et al., 1994). cn~,~lis expressed in antennal lobes, mushroom bodies, neurons in the thoracic ganglia and in muscle fibers (Miyazu el a/., 2000). In addition to the C N G channels, there are two other families of ion channels that contain cNMP-binding sites. These are the hyperpolarizationactivated (lh) channels (Ludwig el al., 1998) and the ether-(t-,4o-~o (eag) family of voltage activated potassium channels. Two lh channels have been reported in insects, one in Helioflfis ~,irescens (Krieger et al., 1999) and the other in Drosophiht (Marx er al., 1999). The mammalian lh channels are activated b~

32


hyperpolarizing currents and the presence of either cAMP or c G M P shifts the voltage dependence, making the channel open at more positive voltages (Ludwig e t a ] . , 1998). The Helioth& lh channel has been expressed in Spodoplera cells and was activated by both cAMP and hyperpolarizing current pulses (Krieger el al., 1999). Both insect channels show prominent expression in sensory tissues such as the eye, antennae and auditory organs in Drosophila (Marx el al., 1999) and in antennal olfactory sensory neurons in Heliothis (Krieger el al., 1999). The Drosophila eag family of ion channels contains three members eag,, eag-like K" channel (elk) and sei:ure (sei) (Littleton and Ganetsky, 2000). When expressed in heterologous cells, depolarizing voltages activated the eag channel and cAMP shifted the threshold for activation to more negative voltages (Brfiggeman eta]., 1993). The application of c G M P appeared to have no direct effect, although it is possible that P K G might modulate eag (Brfiggeman el a]., 1993). There are no reported studies on the effects of cAMP or c G M P on the elk or sei channels.

3.2

PHYSIOLOGICAL FUNCTIONS

The two systems where c G M P regulation and function have been most extensively studied are probably mammalian photoreceptors (Stryer, 1986) and vascular smooth muscle (Lincoln, 1989). In vertebrate photoreceptors, c G M P is the primary transduction signal, c G M P binds to and opens cGMPgated channels maintaining the dark current. Subsequently, light-triggered c G M P hydrolysis causes channel closing and hyperpolarization (Stryer, 1986). Both NO-sensitive soluble GCs and receptor GCs regulate the levels of c G M P in vascular smooth muscle. NO is released from the neighboring endothelial cells that line the blood vessels and activates soluble GC (Bredt and Snyder, 1994), while the atrial natriuretic peptides are released from the heart and regulate receptor GCs (Drewett and Garbers, 1994). The c G M P thus formed activates cGMP-dependent protein kinases, resulting in the phosphorylation of several proteins and smooth muscle relaxation (Lincoln et al., 1994). Both systems act in concert to regulate smooth muscle tone and blood pressure. Neither of these systems has a direct parallel in insects. Insect phototransduction does not involve cGMP, but rather utilizes a light-stimulated phospholipase C and activation of TRP (transient receptor potential) and TRP-like channels (Montell, 1999). The open circulatory system of insects means that there is no parallel to vascular smooth muscle, and there are no reports that c G M P plays a role in modulating the tension of insect skeletal or visceral muscles, although glutamate appears to increase c G M P levels in insect skeletal muscle (Robinson et al., 1982). Nevertheless, there are several examples of physiological processes where there is substantial evidence that c G M P plays a major regulatory role.


3.2.1

33

Sensory physiology

The genetic analysis of phototransduction in Drosophila has described the signal transduction pathways in great detail and shows that lnsP3 formation is the primary signal (Montell, 1999). There is, however, evidence showing cGMP involvement. There are several examples where photoreceptors express genes, which are known to participate in the formation or action of cGMP, that implies a role for cGMP. NO-sensitive soluble GC activity has been detected in locust eyes and immunoreactivity for the ce subunit has been localized to locust photoreceptors, where it has been suggested to play a role in dark adaptation (Jones and Elphick, 1999). Electroretinogram (ERG) recordings from locust eyes showed that cGMP and NO donors increased the response of the photoreceptors to light (Schmactenberg and Bicker, 1999). NO-sensitive GC activity has also been detected in Drosophila photoreceptors during adult development (Gibbs and Truman, 1998), where it has been suggested to play a role in synapse formation (Gibbs and Truman, 1998). Support for this idea has been provided by an analysis of DgceH mutations, which failed to show phototaxis (Gibbs el el/., 2001). ERGs showed that this was not due to a failure of the primary phototransduction cascade but to defects in the post-synaptic responses to light stimuli (Gibbs el al., 2001). The Drosophila cng and ]h channels are also expressed in photoreceptors (Baumann el ell., 1994; Marx el al., 1999). Application of exogenous cGMP induced membrane currents and also enhanced light-induced currents in Drosophila photoreceptors (Bacigalupo et ell., 1995). Similarly, although the primary transduction pathway in insect olfactory neurons does not seem to be mediated by cGMP, the presence of various elements of the cGMP pathway suggests that it is involved in some aspect of olfactory sensation. In most vertebrate olfactory neurons, the primary transduction signal is cAMP, which acts on a CNG channel (Brunet et ell., 1996). A large family of G protein-coupled receptors has been described that are believed to act as the olfactory receptors that couple to adenylyl cyclase (Buck and Axel, 1991). Drosophila also has a large family of G protein-coupled receptors that are expressed in olfactory and gustatory neurons (Clyne el al., 1999; Vosshall et el/., 1999). Pheromone stimulation of silkmoth antennae triggers a rapid and transient increase in lnsP3 (Breer el ell., 1990) and mutations in the Drosophila noq)A gene that encodes a PLC significantly reduced olfactory signaling (Riesgo-Escovar et a/., 1995). These studies suggest that the primary signal for most olfactory stimuli is InsP3. Both soluble and particulate GC enzyme activity have been measured in silkmoth antennae (Ziegelberger et al., 1990), and the Mamtuca receptor-like GC, MsGC-I has been detected in olfactory receptor neurons (Nighorn et al., 2001). In addition, the Drosophila cng, lh and Heliothis lh channels are expressed in antennae (Baumann et ell., 1994; Marx et ell., 1999; Krieger et al., 1999). Pheromone stimulation of silkmoth antennae also triggered an

34


increase in c G M P levels (Ziegelberger et al., 1990), but the time course of increase was much slower than the increase in InsP3, and was suggested to play a role in adaptation. Interestingly, the GC enzyme activity and the pheromone-stimulated c G M P increases measured in silkmoth antennae were detected in whole antennae but not in the olfactory dendrites, providing additional evidence against a primary signal transduction role for c G M P in olfaction (Ziegelberger et a/., 1990). Further evidence for a modulatory role of c G M P in olfactory processing comes from studies on C. ele~ans. A large family of receptor GCs has been described in C. e/egans, many of which are expressed in olfactory neurons (Yu et al., 1997). Two of these, ODR-I and DAF-I 1, are expressed in the chemosensory neuron, AWC, and mutations to either gene show that both are necessary for chemotaxis to all AWC-sensed odorants (Bargmann et al, 1993; Birnby et al., 2000). Deleting the extracellular portion of ODR-I showed that it was not required and hence did not function as the olfactory receptor itself (L'Etoile and Bargmann, 2000). Additional experiments showed that ODR-1 was necessary for odor adaptation and discrimination (L'Etoile and Bargmann, 2000), again suggesting a modulatory role of cGMP in olfactory signaling. In addition to these potential roles in visual and olfactory sensory neurons, c G M P signaling has also been implicated in sensory information processing in primary sensory interneurons and higher order centers (Mfiller, 1996; Nighorn et al., 1998, 2001). Most of the evidence for a role in these processes is the localization of GC and NOS expression to the antennal and optic lobes and central brain structures, although some functional studies have been carried out (see Mfiller, 1996: Bicker, 2001).

3.2.2

Neuronal development

There is accumulating evidence for a role of c G M P in neuronal development, particularly in axonal pathfinding and synapse formation. A wide variety of studies have identified many cell surface and secreted proteins that regulate axonal pathfinding and synapse formation in a wide variety of insects (TessierLavigne and Goodman, 1996). Although several studies have described downstream effectors for some of these proteins (Mueller, 1999L for the most part the signal transduction events that mediate axonal pathfinding are still relatively poorly understood. Nevertheless, evidence from a variety of different experimental systems have shown that c G M P can play an important role in these developmental events. Several studies have shown that NOS, NOsensitive soluble GCs and/or NO-stimulated c G M P immunoreactivity are localized to neurons at times in development when they are growing or nearing their targets. These studies include locust motor neurons (Truman et al., 1996; Ball and Truman, 1998) and antennal pioneer neurons (Seidel and Bicker, 2000), Drosophila photoreceptors (Gibbs and Truman, 1998) and Manduca


35

enteric (Wright et al., 1998) and olfactory neurons (Gibson and Nighorn, 2000). There have also been studies where investigators have blocked NOS and/or NO-sensitive soluble GC and have shown that these treatments have disrupted outgrowth and patterning. During development of the enteric nervous system of M a n d u c a , neurons migrate along the midgut and then extend processes on to the gut musculature (Copenhaver and Taghert, 1989, 1990). Prior to migration none of the enteric neurons showed NO-stimulated GC activity, but soon after the onset of migration a subset (about 50%) of the neurons showed strong NOstimulated c G M P immunoreactivity (Wright e t a / . , 1998). These cells continued to show a NO-stimulated increase in c G M P as they ended migration, extended axons and elaborated processes on to the gut musculature (Wright et al., 1998). Blocking either NOS or NO-stimulated soluble GC activity with a variety of reagents had no effect on the extent of migration or axonal outgrowth (Wright et al., 1998). However, the extent of the terminal branches formed during the final stages of differentiation was significantly reduced when embryos were treated with NOS and soluble GC inhibitors. In addition, the level of immunoreactivity to the synaptic vesicle protein, synaptotagmin, was also significantly decreased by blocking NOS and soluble GC (Wright et al., 1998). These results suggest that the migration and axon formation of enteric neurons does not require NO-stimulated cGMP, but some aspect of terminal differentiation or synapse formation requires increased c G M P levels (Wright et aL, 1998). A similar situation is seen with the development of photoreceptors in Drosophila. During metamorphosis, the photoreceptor axons grow into the CNS to form a precisely ordered projection pattern (Wolff and Ready, 1993). When the tissue was incubated with NO donors during this time, the photoreceptors' axons showed an increase in cGMP, whereas at earlier and later times in development the sensitivity to NO-donors was not apparent. These results suggested that c G M P was involved in the formation of these patterns (Gibbs and Truman, 1998). These predictions were confirmed by maintaining the developing eyes and CNS in tissue culture. When the tissue was incubated with NOS and soluble GC inhibitors the axonal projection patterns were disrupted, but could be rescued with the addition of exogenous c G M P (Gibbs and Truman, 1998). The axons tended to overgrow their normal termination zones, suggesting that NO-stimulated c G M P was necessary to signal the axons to stop growing and to differentiate and form synapses (Gibbs and Truman, 1998). Surprisingly, when flies with a disrupted gene for the c~ subunit of NO-stimulated soluble GC were examined, no defects were seen in the gross organization of the axonal projection patterns (Gibbs et al., 2001). Closer examination of these flies, however, showed defects in their phototaxis behavior and electroretinograms revealed deficits in photoreceptor synaptic transmission (Gibbs et a/., 2001). Thus, in a similar manner to the development of M a n d u c a enteric neurons, NO-stimulated c G M P is not

36


necessary for axonal outgrowth, but is required for termination of growth and synapse development. There is also evidence that the activity of the Drosophila PKG./br mediates neuronal outgrowth at the neuromuscular junction (NMJ). As described previously (section 3.1.1) there are two naturally occurring alleles of the jbr gene that express different levels of P K G (Osborne et al., 1997). When the NMJ of flies bearing the different alleles were examined they showed significantly different branching patterns and projections, suggesting that PKG played a role in their formation (Renger et al., 1999). Here, as with the photoreceptor axons, the alleles with the lower levels of P K G activity showed additional branching of motor neuron axons (Renger et al., 1999). It is possible that again, the reduced levels of a c G M P signaling system prevent the normal terminal differentiation and formation of synapses. These effects might not be direct, however, as different levels of spontaneous electrical activity and evoked transmitter release were also observed and activity patterns are well known to influence neuronal branching and outgrowth (e.g. Budnik et al., 1990). A contrasting situation exists in the developing antennae of locusts. In this tissue, axonal outgrowth rather than the terminal differentiation of the developing olfactory receptor neurons is inhibited by the addition of NOS and soluble GC blockers (Seidel and Bicker, 2000). Thus, there appears to be more than one role for c G M P in developing neurons of insects. Future studies are likely to reveal additional roles and it is especially intriguing to note that the presence of c G M P in the nuclei of developing neurons has been described, suggesting a role for c G M P in the regulation ofgene expression (Truman el al., 1996; Seidel and Bicker, 2000). Although these studies strongly support a role for c G M P in neuronal development, there is no information on the nature of the signals that activate the N O / c G M P pathway. A possible candidate for this has, however, emerged from studies in vertebrate neurons. The secreted protein, semaphorin III, induces the directed growth of growth cones in cultured Xenopus spinal neurons. Simultaneous application of c G M P analogs with semaphorin 1II will reverse the direction of this growth (Song et al., 1998). Semaphorin III also acts as a chemoattractant for dendrites of pyramidal neurons of mice, yet acts as a repulsive agent for the axons of the same neurons. Interestingly, there is a differential distribution of soluble GC in these cells, with a higher concentration in dendrites compared to axons (Polleux el al., 2000). This suggested that a differential distribution of c G M P led to the different responses - a model supported by the finding that blocking soluble GC activity prevented dendritic outgrowth but had no effect on axonal outgrowth (Polleux et al., 2000). Semaphorin family members have been identified in grasshoppers and Drosophila, where they are also essential in axonal pathfinding and the formation of correct synaptic junctions between motor neurons and muscles


37

(Kolodkin et al., 1997: Winberg el al., 1998). Thus the possibility exists that in insects the effects of semaphorin are also mediated by cGMP. 3.2.3

Ecd~,sis"

Probably one of the best studied roles for c G M P in insects is the action of eclosion hormone and the regulation of ecdysis behavior. At the end of each molt, insects need to escape from the cuticle of the previous instar. This is accomplished by a stereotyped behavioral sequence known as ecdysis, or eclosion for adult ecdysis (Reynolds, 1980). Initiation of ecdysis and the preparatory behavior, pre-ecdysis, involves a positive feedback loop between two peptides, eclosion hormone (EH) and ecdysis-triggering hormone (ETH) (Zitnan el al., 1996, 1999: Ewer el al., 1997: Gammie and Truman, 1997a, 1999: Zitnan and Adams, 2000). EH is a 62 amino acid peptide that is located in two pairs of neurons in the brain (Truman and Copenhaver~ 1989) and is released both centrally into the neuropil of the ventral nerve cord (VNC) and peripherally into the circulatory system (Hewes and Truman, 1991). Early studies on the role of EH in triggering ecdysis behavior showed that the action of EH was mediated by an increase in the levels o f c G M P in the VNC (Truman et al., 1979, Morton and Truman, 1985). Subsequent immunocytochelnical studies identified three specific cell populations that increase their c G M P levels in response to EH. Circulating EH acts on peripherally located epitracheal glands (EG). The epitracheal glands are located near each of the 18 spiracles and each contains four cells (Zitnan el al., 1996, 1999: Klein el al., 1999). The glands respond to the action of EH with an increase in c G M P (Ewer et al., 1997: Kingan et el/., 1997) and one of the four cells, the Inka cell, releases its content of ETH (Zitnan el ell., 1996: Kingan el al., 1997). ETH then forms a positive feedback loop by acting on the EH-containing cells in the brain to trigger further release of EH (Ewer et al., 1997). Centrally released EH acts on a population of about 50 neurons in the ventral ganglia that release another neuropeptide, crustacean cardioactive peptide (CCAP) (Ewer el al., 1994: Gammie and Truman, 1997a). The CCAP cells also show an increase in c G M P and the EH/cGMP-triggered release of CCAP is believed to directly activate the ecdysis motor program (Gammie and Truman, 1997a, 1999). A third EH target has been identified that does not appear to be directly involved in ecdysis behavior. Isolated VNCs from prepupal M a m l u c a responded to EH with a large increase in c G M P (Morton and Giunta, 1992) that was restricted to the transverse nerves of each abdominal ganglion (Morton, 1996). This EH-stimulated c G M P increase was localized to a population of intrinsic cells located in the posterior region of the transverse nerve, named the sub-transverse nerve region (STNR) (Hesterlee and Morton, 2000). The STNR cells are believed to develop into the ventral diaphragm muscles that ultimately lie dorsal to the VNC in the adult (Champlin el a/., 1999: Hesterlee and Morton, 2000).

38


A series of studies showed that the EH-stimulated c G M P increase occurs via a novel mechanism that does not utilize an NO-sensitive soluble GC, or a receptor GC, but instead appears to activate an NO-insensitive soluble GC (reviewed in Morton and Simpson, 2001). The primary evidence for this was that EH stimulated a c G M P increase in intact tissue, but failed to do so in tissue homogenates (Morton and Giunta, 1992). This property is a characteristic of pathways that utilize a soluble GC, as ligand-stimulated receptor GC activity can often be demonstrated in cell free preparations. The EH-stimulated c G M P increase in VNCs was not blocked by a variety of NOS inhibitors and NO donors did not mimic the action of EH in any of the EH target cells described above (Ewer et al., 1994; Morton, 1996; Kingan et al., 1997). This demonstrated that the GC present in the target cells was not an NO-sensitive soluble GC. These findings triggered the search for novel GCs that led to the identification of MsGC-I and MsGC-fl3, both potential candidates for the EHstimulated GC as they are NO-insensitive soluble GCs (see section 2.1.3). MsGC-I is not expressed in any of the target cells, ruling it out as a candidate (Simpson et al., 1999). Recent data have shown, however, that MsGC-/33 is expressed in both the STNR and the Inka cells providing circumstantial evidence that MsGC-fl3 is activated by EH in these cells (Morton, 2000; Morton and Simpson, 2001). A major question that remains to be resolved is the mechanism that is used to activate MsGC-fl3. Studies using M a n d u c a VNCs showed that EH stimulated an increase in lnsP3 levels (Morton and Simpson, 1995), and a variety of inhibitors of lipid metabolism blocked the EH-stimulated increase in c G M P (Morton and Giunta, 1992; Morton and Simpson, 1995). These studies suggested that production of a lipid intermediate was necessary for c G M P production, and the most likely pathway appears to involve activation of protein kinase C that then leads to the activation of MsGC-fl3 (see Morton and Simpson, 2001). As MsGC-fl3 is not expressed in the CCAP ceils, an alternative pathway is likely to be used in these ceils. Whether this involves another NO-insensitive soluble GC or whether they express an EH-stimulated receptor GC is not known. The physiological events downstream of the EH-stimulated increase in c G M P are cell specific. In the Inka cell of the epitracheal glands, the EHstimulated c G M P increase mediates the release of ETH (Kingan et a/., 1997). The release of ETH also involves increases in cytoplasmic calcium levels. Studies on isolated epitracheal glands showed that release of intracellular stores of calcium in the lnka cells using thapsigargin stimulated ETH release and EH triggered an increase in intracellular calcium levels (Kingan et al., 2001). Incubation of epitracheal glands with 5raM cobalt in the absence of extracellular calcium had no effect on the EH-stimulated release of ETH~ suggesting that release did not require extracellular calcium (Kingan et al., 1997). It has been suggested that EH triggers a parallel increase in both c G M P and calcium in the Inka cells, which act in concert to trigger ETH release (Kingan et al., 2001). Protein phosphorylation acts downstream of


39

the c G M P accumulation to mediate EH-stimulated ETH release. Staurosporine, a broad spectrum protein kinase inhibitor, blocked ETH release but had no effect on EH-stimulated c G M P levels (Kingan et al., 2001). In addition, the protein phosphatase inhibitor calyculin A, potentiated both basal and EHstimulated ETH release (Kingan et al., 2001). These results suggest that activation of a cGMP-dependent protein kinase (PKG) is a likely mediator of ETH release (Kingan et al., 2001). EH also appears to act on the CCAP cells as a peptide-releasing hormone, stimulating the release of CCAP, which then triggers the ecdysis motor program (Gammie and Truman, 1999). Although there is considerable circumstantial evidence that EH acts directly on the CCAP cells, stimulating CCAP release via an increase in c G M P levels, there is, as yet, little direct proof for this pathway. Electrical stimulation of the VM cells (to stimulate EH release) and application of EH to desheathed abdominal ganglia cause c G M P increases in the CCAP cells and initiation of the ecdysis motor program (Gammie and Truman, 1999; Zitnan and Adams, 2000). Application of CCAP to desheathed ganglia also triggers the ecdysis motor program (Gammie and Truman, 1997a). Ecdysis triggered with the application of CCAP is initiated with a shorter latency than with EH and there is no corresponding increase in c G M P indicating that EH action is upstream of both the c G M P increase and CCAP release (Gammie and Truman, 1999). However, the ability of EH to induce the ecdysis motor program varies between individual preparations, and this variability has been interpreted to indicate that the effects of EH on the CCAP cells are not direct (Zitnan and Adams, 2000). Another explanation is that multiple inputs on to the CCAP cells are necessary for CCAP release, one of which is EH. Evidence to support this comes from Drosophila, where the absence of EH does not prevent ecdysis in all animals, although it does result in a less robust expression of the behavior (McNabb et al., 1997). This is consistent with the presence of multiple pathways activating the CCAP cells. As EH increases c G M P by direct action on the epitracheal glands and the STNR cells, it seems likely that it also acts directly on the CCAP cells to increase c G M P levels. The effect of increased c G M P levels in one of the CCAP cells, cell 27, has also been investigated (Gammie and Truman, 1997b). At times in development prior to the endogenous increase in cGMP associated with ecdysis, cell 27 showed a higher threshold for firing action potentials compared with times when c G M P was elevated. In addition, there were no spontaneous action potentials in the absence of an increase in cGMP, whereas, after c G M P levels had risen, spontaneous action potentials at frequencies of up to 2 Hz were recorded (Gammie and Truman, 1997b). Application of the cell-permeant analog, 8-bromo-cGMP mimicked the decrease in action potential threshold (Gammie and Truman, 1997b). Pharmacological and ion-substitution experiments suggested that this cGMP-induced reduction in threshold was mediated by enhancing an inward calcium current (Gammie and Truman, 1997b). These

40


electrophysiological changes could then lead to CCAP release, either directly or by enhancing the efficacy of additional synaptic inputs. The STNR cells of the abdominal transverse nerves also respond to EH with an increase in cGMP. The physiological significance of this increase is, however, unclear. The STNR cells are present throughout larval life and go through a period of enhanced proliferation during pupal development (Champlin et al., 1999: Hesterlee and Morton, 2000). The cells also become sensitive to EH during pupal development. They show no increase in c G M P in response to EH at larval ecdysis or during the final larval instar, but first show an EH-stimulated c G M P increase soon after the wandering stage at the onset of pupal development (Hesterlee, 1999). Although it has not been demonstrated directly, the STNR cells are likely targets for circulating EH at pupal ecdysis (Morton, 1996, 1997). Soon after ecdysis, the cells spread out from their tightly grouped position along the transverse nerve to become a thin sheet of cells covering each abdominal ganglion (Hesterlee and Morton, 2000) and subsequently develop into the ventral diaphragm muscles in the adult (Champlin el al., 1999; Hesterlee and Morton, 2000). Several possible roles for the EH-stimulated c G M P increase have been suggested, such as triggering the migration or initiating the differentiation of the cells, but as yet there is no evidence to support these possible actions (Hesterlee and Morton, 2000). Although the physiological significance of the EH-stimulated c G M P increase in the STNR cells is unknown, there is circumstantial evidence that the EGPs (described in section 3.1.1 ), proteins that are phosphorylated in response to EH and c G M P are located in the STNR cells. The most compelling evidence for this comes from experiments with isolated VNCs exposed to EH. Immunocytochemistry showed that in these preparations EH stimulated a c G M P increase only in the STNR cells (Morton, 1996). No increase was seen in the CCAP (crustacean cardioactive peptide) cells, as circulating or bath-applied EH cannot cross the blood brain barrier (Gammie and Truman, 1999). Nevertheless, incubation of isolated VNCs with EH stimulated the phosphorylation of the EGPs in a cGMP-dependent manner (Morton and Truman, 1988a), suggesting that the EGPs are localized to the STNR cells. Thus, the E H / c G M P pathway plays a central role in the physiology of ecdysis. In the Inka and CCAP cells it leads to the release of additional peptides and activation of the ecdysis motor program. It is likely that EH also functions as a master coordinating hormone regulating additional physiological events associated with ecdysis. The programmed degeneration of the intersegmental muscles of the silk moth Antheraea polyphemus lbllowing adult ecdysis is thought to be triggered by the direct action of EH via an increase in c G M P (Schwartz and Truman, 1982, 1984). The Verson's glands in Manduca release their contents at ecdysis a process that is prevented when peripherally released EH is blocked (Hewes and Truman, 1991) although it is not known whether this is the result of the direct action of EH or whether it is mediated by cGMP. Studies of eclosion in Drosophila revealed that an increase in c G M P


41

was occasionally seen in tracheae that correlated with the release of EH (Baker et al., 1999). These c G M P increases were absent in animals that lacked EH, suggesting that tracheae are also EH targets (Baker et al., 1999). Many of the animals that lacked EH were unable to correctly fill their tracheae with air following ecdysis, suggesting a role for the E H / c G M P pathway in this process (Baker el al., 1999). The action of EH on the STNR cells is also likely to be part of this repertoire of physiological events that are associated with ecdysis. 3.2.4

k~)od-seareh behavior

A genetic analysis of food searching strategies in Drosophila has shown that c G M P plays a central role in this behavior (Sokolowski, 1998). Two naturally occurring alleles of the Jbraging (/or) gene exhibit different behaviors. In the presence of food, flies with the sitter allele stayed relatively stationary and remained in a single patch of food. By contrast, flies with the rover allele continued to forage for additional food and moved between patches of food (Sokolowski, 1980). In the absence of food, however, both alleles showed similar levels of locomotory activity, showing that sitter flies were not simply more sluggish and that the behavioral differences were foraging specific. Several lines of evidence showed that the jbr gene codes for a P K G (Osborne et al., 1997). Both the naturally occurring alleles and additional mutations in t h e j b r gene mapped close to a region that contains a previously identified gene for PKG (Osborne et al., 1997). The levels of total P K G enzyme activity were measured in these strains and showed that in flies with the naturally occurring sitter alleles, and in jbr mutations that exhibited a sitter phenotype, there was a slightly reduced, but significantly lower level of P K G present (Osborne et al., 1997). Flies containing a P element that was located in, and disrupted the open reading frame of, one of the jbr PKG transcripts, showed a sitter phenotype, and excision of the P element caused flies to revert back to the rover phenotype (Osborne et al.. 1997). Further proof that ./br encodes a P K G came fi'om overexpression of a cDNA for the PKG in sitter flies. This manipulation caused an increase in PKG levels and a rover phenotype (Osborne et al., 1997). Northern and western blots showed that both RNA and protein levels for t h e / b r P K G were lower in sitters than rovers (Osborne et al., 1997). A preliminary description of the expression patterns o f l o t showed widespread distribution in olfactory, gustatory, gut and brain tissues (Sokolowski, 1998). This leaves open several possible levels of organization that could be modulated by the c G M P / P K G pathway resulting in the behavioral differences. Expression o f . / o r in chemosensory tissues suggests that different levels of chemosensation associated with food might generate the different foraging strategies. The finding that the behavioral differences are only apparent in the absence of food supported this idea and, in addition, many different studies have provided evidence for a role of c G M P in modulating olfactory pathways

42


(see section 3.2.1). Behavioral differences associated with the presence or absence of food could also be activated by signals originating in the gut, and the presence of Jbr in gut tissue suggests that this is also a possibility. Expression of Jbr in the central regions of the brain, and the finding that differences of PKG activity could be detected in the heads of sitter and rover strains, suggest that differences in the central processing of information are also likely to represent a large component of the behavioral phenotype. A series of physiological studies are consistent with this suggestion. Cultured giant neurons from strains that showed the sitter phenotype (both natural alleles and induced mutations) showed hyperexcitability and reduced voltagedependent potassium currents compared with animals with rover phenotypes (Renger et al., 1999). Similarly, recordings from the neuromuscular junction showed that sitter strains exhibited higher levels of spontaneous activity and larger evoked post-synaptic potentials as compared with rovers (Renger et al., 1999). Thus it is likely that cGMP/PKG-mediated modulation of the nervous system is necessary at several levels of organization to generate the highly specific phenotypical differences in the two strains of flies. Clearly, many more studies are needed to gain a clearer understanding of this complex behavior and the role that cGMP signaling plays in its production. 3.2.5 Malpighian tubule regulation The regulation of fluid secretion from Malpighian tubules is another system for which there is good evidence for a central role of cGMP. Malpighian tubules are the principal excretory organ in insects, responsible for the production of primary urine (Chapman, 1998). In most insects, potassium is actively pumped into the tubules, and other solutes and water then follow passively. Resorption of ions, amino acids, water, sugars and other necessary components is then accomplished in the Malpighian tubules themselves or in the ileum and rectum (Chapman, 1998). The rates of both excretion and resorption are regulated by a variety of circulating hormones (Chapman, 1998). The majority of the studies that implicate cGMP as a regulator of Malpighian tubule fluid secretion have been carried out in Drosophila. RT PCR experiments have shown that Drosophila Malpighian tubules express the PKGs DG1 (Pkg21D) and DG2 (/br) (Dowet al., 1994), NOS (Davies et al., 1997) and the cGMP-gated ion channel cn~, (MacPherson et al., 2001). Experiments on isolated tubules show that both NO donors and exogenous cGMP stimulate fluid secretion and zaprinast, an inhibitor of cGMP-specific PDEs, and okadaic acid, a protein phosphatase inhibitor, potentiates the NOstimulated and cGMP-stimulated fluid secretion (Dowet al., 1994). At least two neuropeptides also stimulate fluid secretion: leucokinin (Terhzaz et al., 1999) and cardioacceleratory peptide 2b (CAP2b) (Davies et al., 1997). Both leucokinin and CAP2b stimulate an increase in intracellular calcium levels, although the effects of this increase differ with the two peptides. Leucokinin


43

does not affect c G M P levels and stimulates an increase in chloride permeability, probably acting on the stellate cells of the tubule (O'Donnell el al., 1996; MacPherson, 2001). By contrast, CAP2b acts on the principal cells of the tubule to increase the activity of an apical vacuolar type H +-ATPase (VATPase) that stimulates cation transport across the tubule epithelium (O'Donnell el al., 1996). The increase in intracellular calcium triggered by CAP2b stimulates NOS, which in turn stimulates an increase in the levels of c G M P (Davies el al., 1997). The increase in c G M P probably has two effects. The major effect is to stimulate P K G activity, which activates the V-ATPase (O'Donnell et al., 1996) and there is evidence that cGMP also activates a cation-selective channel (possible the cng channel) to cause a longer lasting increase in intracellular calcium (MacPherson el a/., 2001). It is not clear how widespread this role of c G M P is in other insects. In Rhodnius prolixus Malpighian tubules, CAP2b also stimulates an increase in c G M P levels, but both CAP2b and c G M P inhibit rather than stimulate fluid secretion (Quinlan et al., 1997). Blood-feeding insects face special problems in urine production, as they need to rapidly excrete large quantities of water (Chapman, 1998). The anti-diuretic effects o f c G M P may therefore be a special case, but studies on the role of c G M P in Malpighian tubules of additional insects will be needed to determine whether stimulation or inhibition of fluid secretion is the more general action.

4

Concluding remarks

In this review we have described our current understanding of c G M P regulation and function in insects. In addition to reviewing published reports of the c G M P signaling cascade components in insects, we have summarized several of the major physiological systems that utilize cGMP. These studies highlight the diverse roles that c G M P plays in cellular physiology. There have been relatively few molecular studies that have characterized elements of the c G M P cascade in insects, but the availability of the Drosophila genome sequence now enables us to identify and predict the properties of all these components in a single organism. This analysis has revealed a wealth of genes, some of which have previously been described, but most have yet to be characterized. The list of genes that are predicted to code for components of the c G M P cascade include five receptor GCs, two receptor-like GCs, five soluble GC subunits, a possible GC-activating peptidc, five possible GCAPlike proteins, seven PDEs (of which five are predicted to hydrolyze c G M P and two might be regulated by cGMP), three PKGs, and a total of eight ion channels with cNMP-binding sites, of which at least two could be regulated by cGMP. Remarkably, there have been few genetic studies that describe the outcome of mutations in these genes. The systems that have been studied in this manner have shown profound effects: alterations in food search behavior and

44


defective synaptic connections, suggesting that this will be a fertile area for future investigations.

Acknowledgements We wish to thank Dr Alan Nighorn of the University of Arizona for providing unpublished sequence information, and Kristofor Langlais and Steve Matsumoto for comments on the manuscript. This work was supported by N I H grant NS29740.

Note added in proof A recent report (Tanoue et al., 2001) has described the cloning of a receptor G C from the silkmoth B o m b y x mori, The sequence of the extracellular domain of this GC, named BmGC-I, is almost 30'70 identical to that of mammalian atrial natriuretic peptide (ANP) receptors, including 100% conservation of the cysteine residues. This suggests that in B o m b y x , there may be ANP-like peptides that activate this GC. BmGC-I is widely expressed, including expression in olfactory sensory neurons in the antennae, glomeruli of the antennal lobes, thoracic ganglia, flight muscles, midgut and Malpighian tubules. Another recent report showed that the expression of BmGC-I in flight muscle is under circadian control (Tanoue and Nishioka, 2001). Tanoue, S., Sumida, S., Suetsugu, T., Endo, Y. and Nishioka, T. (2001). Identification of a receptor type guanylyl cyclase in the antennal lobe and antennal sensory neurons of the silkmoth, Bombvx mori. bisect. Biochem. Mol. Biol. 31, 971 979. Tanoue, S. and Nishioka, T. (2001). A receptor-type guanylyl cyclase expression is regulated under circadian clock in peripheral tissues of the silk moth. Light-induced shifting of the expression rhythm and correlation with eclosion. J. Biol. Chem. 276, 46 765-46 769.

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Neurotransmitter Transporters in the Insect Nervous System Stanley Caveney a and B. Cameron Donly b ~Department of Biology, University of Western Ontario, London, Ontario N6A 5B7, Canada ~Southern Crop Protection and Food Research Centre, Agriculture and Agri-Food Canada, London, Ontario, Canada N5V 4T3

I

Introduction 56 1.1 Background 56 1.2 Molecular chemistry of insect neurons 56 1.3 Neurotransmitter uptake and vesicular storage 59 1.4 Scope of the review 61 2 Excitatory amino acid transporters 61 2.1 Net+:K+-dependent glutamate transporters 61 ") "~ Net ,K -dependent aspartate transporter 77 3 Na+/CI -dependent GABA and monoamine transporters 1 78 3.1 GABA transporters 79 3.2 Serotonin transporters 90 3.3 Dopamine transporters 99 3.4 Octopamine transporters 106 3.5 Orphan transporters 111 4 Na~:(71 -dependent transporters II 114 4.1 Choline transporters 114 5 Other Na ~-dependent transporters 121 5.1 Histamine transporter 121 6 Putative neurotransmitter transporters 123 6.1 Glycine transporters 124 6.2 Taurine transporters 124 7 Applications to insect control 125 7.1 Relevance of insect neurochemistry to pest control 125 7.2 Neurotransmitter transporters as new' targets for insect control 7.3 Future directions 127 7.4 Post-genomic prospects t\~r insect physiology research 128 Acknowledgements 129 References 129

125

Note: For the sake of consistency' this review uses a three-letter designation for neurotransmitter transporters described fiom invertebrates and a one-letter designation for those described from vertebrates. ADV,%N('t

S IN INSF(T

1%13N i i- I l 02422~) - X

I%tYSIOI O(iY

VOI.

2~)

( op ~i~ht ,

2(1()2 I:/,~e~lcJ ~,¢i¢lt~c l.td

I / / 117]lt~ ~4 ~c/iJodm tlotl lit ~m i h~lm ~'s~,/vl,I

56

1 I.l

S. CAVENEY AND B. C. DONLY

Introduction BACKGROUND

Neurotransmitters are chemical signals released from neurons at specialized synapses by exocytosis (Pennetta et al., 1999; Prokop, 1999). These neurotransmitter molecules diffuse within the synaptic space and bind to specific neurotransmitter receptors on the membranes of post-synaptic neurons, muscle cells, peripheral effector cells, glial cells surrounding the synapse, or even on the surface of the neuron that released the neurotransmitter (autoreceptors). The neurotransmitter receptor interactions involved normally change the behaviour of a target cell by altering its membrane potential or second messenger signalling pathways. This signalling event is rapidly terminated because the neurotransmitter molecules disappear quickly from the synaptic space, either by diffusion or by local uptake (as intact or partially degraded molecules) by neuroglial cells. A second class of cell surface proteins at the synapse, neurotransmitter transporters (NTTs), are responsible for the removal of neurotransmitter from the synaptic space (Fig. 1). These NTTs may be located on the surface of the pre- or post-synaptic neurons, or on the surface of glial ceils that surround the synapse. The pattern of rapid transmitter "reuptake' following transient transmitter release is an essential feature of chemical neurotransmission. Transmitter molecules taken up by axon terminals are concentrated and stored in synaptic vesicles for future use. The number of neuroactive chemicals taken up by neurons and stored in synaptic vesicles in insects is known to be rather limited. Fewer than ten amines and amino acids appear to fit the stringent criteria that define authentic neurotransmitters (Callec, 1985; Burrows, 1996), implying that only a small number of membrane transport systems may be involved in neurotransmitter recycling at the neural synapse. 1.2

MOLECULAR CHEMISTRY OF INSECT NEURONS

Formerly neurons were classified on the basis of their functional anatomy (sensory neurons, interneurons, motor neurons, region of brain, etc.) and mode of action (excitatory neurons, inhibitory neurons). These epithets reveal nothing about neuronal molecular chemistry, however. In modern times, a neuron is defined by the specific neurotransmitter molecule it stores, releases into the synaptic space and subsequently removes from it (for reviews, see Evans, 1980; Restivo and White, 1990; Buchner, 1991, Burrows, 1996: Osborne, 1996). The rate-limiting enzymes involved in the synthesis and degradation of particular neurotransmitters constitute a set of neuronal 'hallmark' proteins (Buchner, 199l; Lundell and Hirsch, 1994b). Implicit in this terminology is the notion that axon terminals contain plasma membrane transporters that return (or supply) specific neurotransmitters to the axoplasm for re-use.

NEUROTRANSMITTER TRANSPORTERS

57

Presynapse I-

GABA

~:~ Monoamines

H+

onoamines

Na+/CINa ÷

O

GI

Na+ Glu Na+/Cl -

:3K+

O

Glial cell FIG. 1 Neurotransmitter transport at the insect synapse. The diagram is a composite showing known families of Na+-dependent transport proteins in the plasma membrane of the presynaptic axon (left) and glial cells (right), and in the membrane of synaptic vesicles in the axon terminal (top left). Neurotransmitters are actively transported across the plasma and vesicular membrane against a concentration gradient. High-affinity Na+/K+-dependent excitatory amino acid transporters (EAATs, shown at O) transport the neuroactive amino acids glutamate and aspartate across the plasma membranes in glutamatergic neurons or glial cells (as shown). The inhibitory neurotransmitter GABA and several monoamine neurotransmitters (octopamine (OA), dopamine (DA) and serotonin (5hydroxytryptamine, or 5-HT)) are taken up selectively by cells in the CNS through the activity of transporters belonging to one family of N a ' / C 1 - d e p e n d e n t transporters. GABA transporters (GATs O) are found in the plasma membrane of GABA-ergic neurons and glial cells. Monoamine transporters (OATs, DATs, SERTs) are excusively neuronal in distribution (O). Each neuron selectively expresses the monoamine transporter taking up the specific monoamine neurotransmitter that the neuron releases into the synaptic space. Choline, the precursor of acetylcholine, is taken up by chotinergic neurons that express a transporter belonging to an unrelated family of Na +/C1 -dependent transporters (0). Synaptic vesicles further concentrate and sequester neurotransmitters within the axon terminal. In aminergic neurons, this involves the activity of vesicular monoamine transporters (vMATs O), vesicular acetylcholine transporters (vAchTs), vesicular excitatory amino acid transporters (vEAATs) and vesicular inhibitory amino acid transporters (vlAAT). Vesicular neurotransmitter transport is driven by transmembrane H ~ gradients (vMATs) and/ or H--dependent electrochemical gradients (vEAATs). The plasma and vesicular membranes of individual neurons normally contain transporters |\)r a single type:' class of neurotransmitter.

58


Consequently, the high-affinity and Na+-dependent transporters involved are named after the neurotransmitter substrates they selectively transport (Table 1). On the other hand, neurotransmitter receptors, although named for the neurochemical they bind (or after an analogue of the natural transmitter, or an inhibitor of the receptor), are proteins primarily associated with the postsynaptic cells (although presynaptic autoreceptors modulate neurotransmitter release by some insect neurons; see Howell and Evans (1998). The specific proteins involved in neurotransmitter synthesis and transport at the nerve terminal that assign each neuron in the arthropod central nervous system its unique molecular identity are as follows. For cholinergic neurons. the hallmark proteins are the enzyme choline acetyltransferase (CHAT; Buchner et al., 1986; Kitamoto el al., 1998) and the choline transporter (CHT; Wang et al., 2001). Histaminergic neurons in the arthropod retina are identified by the presence of histidine decarboxylase (HDC: Maxwell el al., 1978; Elias and Evans, 1983b; Burg et al., 1993) and of the histamine transporter (HAT; Morgan ez al., 1999). Tyramine fi-hydroxylase (TflH; Monastirioti et al., 1996; Lehman et al.. 2000b), and the octopamine transporter (OAT; Malutan el al., 2002) together act as hallmark proteins for insect octopaminergic neurons. (Trill is the homologue of dopamine fl-hydroxylase that catalyses nor-epinephrine synthesis in mammals). For dopaminergic neurons in insects, as well as other animals, tyrosine hydroxylase (TH; Neckameyer and Quinn, 1989; Lundell and Hirsch, 1994b; Granholm el ell., 1995) and the dopamine transporter (DAT; P6rzgen el ell., 2001) are marker proteins. (DOPA decarboxylase (DDC) is not an unambiguous marker for

TABLE 1 Neurotransmitter transporters as key marker proteins for the different types of neurons in insects Neuron type Cholinergic Serotonergic

Dopaminergic Octopaminergic Histaminergic GABAergic Glutamatergic

Enzymes involved in neurotransmitter synthesis

Na+-dependent high-affinity neurotransmitter transporter

Choline acetyltransferase (CAT) Tryptophan hydroxylase (TPH), Aromatic amino acid decarboxylase (AADC or DDC) Tyrosine hydroxylase (TH), Dopa decarboxylase (DDC) Tyramine hydroxylase (TflH) Histidine decarboxylase (HDC) Glutamic acid decarboxylase (GAD) Glutaminase~' Amino acid transferases ~'

CHT SERT

~ConslittHiveenzymesin rnosl insect cells

DAT OAT HAT GAT EAAT


59

dopaminergic neurons since, as an aromatic amino acid decarboxylase (AADC), it catalyses the final decarboxylation step in both dopamine synthesis (Budnik and White, 1988; Kostal et al., 1998) and serotonin synthesis (Buchner, 1991; Lundell and Hirsch, 1994b) in Drosophila). In serotonergic neurons, tryptophan undergoes a two-step conversion to 5-hydroxytryptamine (5-HT) or serotonin, the first step being catalysed by tryptophan hydroxylase (TPH) (Wright, 1987; Budnik and White, 1988; Lundell and Hirsch, 1994b) and the second by DDC. In addition to TPH, the serotonin transporter (SERT: Corey et al., 1994b; Demchyshyn et al., 1994) is a hallmark protein of serotonergic neurons. The situation is less straightforward for amino acid transporters since many have non-neuronal distributions as well. GABA-ergic neurons in insects are defined by the presence of glutamate decarboxylase (GAD: Baxter and Torralba, 1975; Breer et al., 1989; Jackson et al., 1990) involved in converting glutamate to GABA, but not necessarily by the presence of high-affinity GABA transporters, which occur in glial cells. Because L-glutamate and L-aspartate are amino acids needed in intermediary metabolism and protein synthesis in many types of insect cells, it is not possible to identify glutamatergic neurons solely on the quality of specific enzymes involved in the synthesis or degradation of these amino acids, such as glutaminase and glutamine synthetase, or on the presence of high-affinity glutamate transporters (EAATs). (Glutarnine synthetase and tyrosine decarboxylase activity is found in many tissues in insects, including glial cells in the central nervous system (CNS).) 1.3

N H J R O T R A N S M I T T E R U P T A K E A N D VESICULAR STORAGE

To establish that a candidate molecule is a traditional neurotransmitter, two of the several criteria that must be satisfied are: (1) that a mechanism must exist to terminate the receptor-mediated actions of the putative transmitter following its presumed release into the synapse and (2) that the putative neurotransmitter must be concentrated within synaptic vesicles in the presynaptic terminal (Callec, 1985). With the notable exception of the neurotransmitter acetylcholine, which is degraded by cholinesterase in the synaptic space, neurotransmitters are cleared directly from the space by selective uptake systems (often called re-uptake systems) in the plasma membranes of neurons and/or glial cells, and then sequestered in synaptic vesicles by vesicular transporter systems. Neurotransmitter uptake into the nerve terminal and its synaptic vesicles is required for the nervous system to function normally (Attwell and Mobbs, 1994). Demonstration of a high-affinity membrane transport system and its corresponding lower-affinity vesicular transport system in a nerve terminal (or at least in a nerve preparation) is strong evidence for a candidate neurotransmitter acting as a local neurotransmitter. The molecular biology of the uptake systems that transfer neurotransmitters from the synaptic space into neurons and glial cells has been characterized

60


during the last decade (for recent general reviews, see Reith, 1997; Amara, 1998; Beckman and Quick, 1998; Krantz et al., 1999; Masson et al., 1999). The proteins involved constitute several families of high-affinity Na +dependent neurotransmitter transporters (NTTs) that help to terminate the postsynaptic action of neurotransmitters released from neurons, and replenish their neurotransmitter content. The plasma membrane NTTs for GABA (GATs I 3), serotonin (SERT), dopamine (DAT), nor-epinephrine (NET), glutamate (EAATs 1 5), glycine (GLYT1 and 2), the NT precursor choline (CHT), as well as proline (PROT) and taurine (TAUT) (both putative NTTs) have been cloned from the vertebrate CNS (Masson et al., 1999). Notably absent from this list is a molecular description of the neuronal transporter for histamine. These NTTs belong to three structurally distinct families of co-transporter proteins, all of which are dependent on transmembrane gradients in K + and/or CI , in addition to Na +, for their normal activity. Following the selective uptake of neurotransmitter (or, in the case of acetylcholine, its precursor choline) into a nerve terminal, a low-affinity transport system located in the vesicle membrane concentrates the neurotransmitter in synaptic vesicles (Fig. 1). These vesicle-membrane carriers are powered by proton or voltage gradients across the vesicular membrane and have protein structures distinct from those of plasma membrane NTTs (Attwell and Mobbs, 1994; Worrall and Williams, 1994; Schuldiner, 1997). Vesicular transporters are generally less selective in their substrate preferences compared to plasma membrane transporters, and make up four functional groups, vesicular excitatory amino acid transporters (vEAATs, for glutamate and possibly aspartate), vesicular inhibitory amino acid transporters (vIAATs, for GABA and glycinc), vesicular monoamine transporters (vMATs, for serotonin, catecholamines, histamine, and presumably in insects, octopamine) and the vesicular acetylcholine transporter (vAChT). Molecular cloning studies indicate that there are two mammalian vMAT subgroups, vMATI and vMAT2, that belong to the same protein family as yAChT, and are distinct from the vlAAT (and likely the vEAAT) protein family (reviewed most recently in Schuldiner, 1997; Masson et al., 1999). A few hundred of the several thousand genes expressed in the adult brain of Drosophila code for the enzymes and proteins involved in the presynaptic synthesis, storage, triggered release, re-uptake and/or regeneration of neurotransmitter chemicals, and for the many pre- and post-synaptic ionotropic and metabotropic receptors to which neurotransmitters bind on release into the synaptic space (Buchner, 1991). Almost all brain-related genes in insects have corresponding homologues in mammals (Buchner, 1991; Pennetta el al., 1999). This genetic conservation underscores the value of Drosophila as a general model in neuroscience. Conversely, advances in mammalian molecular neurobiology during the last decade have also facilitated the identification of many new genes and their products in the insect CNS, Considerable progress has been made during the last 10 years in our understanding of the molecular


61

biology of neurotransmitter transport in the insect nervous system. Many of the cDNAs that encode selective high-affinity neurotransmitter transporters in insects have been cloned and characterized. I .4

SCOPE OF THE REVIEW

This review summarizes the current state of understanding of neurotransmitter transport across the plasma membrane in insect nervous tissue. Although research on neurotransmitter transport in the mammalian CNS has intensified over the last decade, a comprehensive review of neurotransmitter transport in the insect CNS has not been available, although Osborne (1996) presented a brief account. The sections in this review describe the molecular physiology of the various N T T systems in insects. The approach used is necessarily hierarchical and comparative. Much of what we know about the molecular biology of neurotransmitter uptake in the CNS of Dro,vophila and other insects is derived from discoveries in the mammalian CNS. The review concludes with an assessment of the potential of neurotransmitter transporters as selective neural targets for future insect control strategies.

2 2.1

Excitatory amino acid transporters Na ~ K ~ - D E P E N I ) E N T

GLUTAMATETRANSPORTERS

L-glutamate is the principal excitatory neurotransmitter in the vertebrate nervous system (Orrego and Villanueva, 1993; Meldrum, 2000). In insects, L-glutamate acts both as an excitatory and inhibitory neurotransmitter (Usherwood, 1994) and is the most abundant amino acid in the CNS (Pitman, 1985). In orthopteroid insects (locust, cockroach), glutamate activates several types of cation-selective channels (fast-acting quisqualate- and kainatesensitive ionotropic receptors), anion-selective channels (ibotenate-sensitive ionotropic receptors) and G-protein coupled receptors (slower-acting metabotropic receptors) (Usherwood, 1994). Several glutamate receptor subtypes have been cloned from Dro,s'ol~hila, and analysis of the FlyBase revealed the presence of other members of the ionotropic glutamate receptor superfamily (Littleton and Ganetzky, 2000). In the honeybee brain, glutamate has been detected immunocytochemically in descending interneurons and in visual interneurons (Bicker et al., 1988). Glutamatergic neurons may be involved in the processing of long-term olfactory memory in the mushroom bodies in the honeybee brain (Maleszka el al., 2000). L-glutamate acts as an inhibitory transmitter at extrajunctional receptors in the locust CNS (Giles and Usherwood, 1985: Wafford and Sattelle, 1989). The ibotenate-selective L-glutamatereceptor/Cl- channels present in insect neurons (Raymond et ell., 2000) are related in molecular structure to GABA-gated C1 channels and not to

62


excitatory quisqualate-sensitive glutamate-gated Na + channels (Lummis et al., 1990; Usherwood, 1994; Osborne, 1996). e-glutamate is also the principal excitatory neurotransmitter at the arthropod neuromuscular junction (Usherwood, 1994; Burrows, 1996). Excitatory glutamatergic motor neurons have been shown, for instance, to innervate skeletal muscle in lobster (Takeuchi and Takeuchi, 1964), Drosophila (Jan and Jan, 1976; Johansen et al., 1989) and locust (Usherwood, 1994; Burrows, 1996), as well as hindgut visceral muscle in Leucophaea (Cook and Holman, 1979). Several hundred neurons, presumably mostly motor neurons, display glutamate-like immunoreactivity in locust thoracic and abdominal ganglia (Watson and Seymour-Laurent, 1993). The excitatory neurotransmitter status of other amino di-acids, such as k-aspartate, L-cysteate and L-cysteine sulphinate, in the insect nervous system is unresolved. These compounds could serve as excitatory neurotransmitters in the mammalian CNS (Griffiths, 1990; Orrego and Villanueva, 1993; Gundersen et al., 1998), and compete with glutamate for Na+-dependent uptake in insect tissues (Caveney et al., 1996). 2.1.1

Background

It has been known for a long time that specific Na+-dependent high-affinity transport plays a key role in inactivating glutamate released at insect neuromuscular junctions (Faeder and Salpeter, 1970; Faeder et al., 1974; Van Marie et al., 1983, 1985). Radiolabelled glutamate uptake was first demonstrated in nerve muscle preparations isolated from Gromphadorhina portentosa (Faeder and Salpeter, 1970), and then in intact abdominal nerve cords from Periplaneta americana (Evans, 1975) and thoracic and abdominal ganglia isolated from the moth Manduca sexta (Kingan and Hishinuma, 1987). The saturable component of glutamate uptake in the CNS was shown to be dependent on external Na +. The sheath (glial) cells are primarily responsible for glutamate uptake by the nerve muscle preparations (Salpeter and Faeder, 1971). Sodium-iondependent and high-affinity uptake of glutamate has also been reported in neurons derived from embryonic cerebral ganglia, but not in embryonic muscle cells, grown in vitro (Bermudez et al., 1988). Synaptosomes prepared from Drosophila also display high-affinity Na+/K+-dependent uptake of L-glutamate and k-aspartate (Ramarao et al., 1987). Employing current mammalian terminology, high-affinity Na+-dependent plasma membrane transporters for L-glutamate and k-aspartate (a putative insect neurotransmitter) are called excitatoo, amino acid transporters, or EAATs, notwithstanding the nonjunctional role of glutamate as an inhibitory neurotransmitter in insects (Usherwood, 1994; Raymond et al., 2000). The less-restrictive acronym GLUT was earlier assigned to cloned facilitated glucose transporters (Thorens, 1996). EAATs are also found in non-neural tissues in mammals (Hediger and Welbourne, 1999; Palacin et al., 1998) where they fulfill a variety of non-neurotransmitter related functions.


63

The molecular biology and pharmacology of EAATs cloned and characterized from the insect nervous system will now be described and compared in light of the wealth of vertebrate information currently available. However, it should be mentioned that several non-excitable insect tissues display Na +dependent uptake of glutamate. These include the epidermis (McLean and Caveney, 1993; Caveney el al., 1996), fat body and hindgut (H. McLean, unpublished) of the beetle T e n e b r i o m o l i t o r and the epidermis of several orthopteroid insects (S. Caveney, unpublished). A role proposed for this non-neuronal uptake is to keep glutamate levels in the haemoplasm below the activation threshold of the neuromuscular synapses (Irving el al,, 1979; McLean and Caveney, 1993; Tomlin el al., 1993). 2.1.2

Slructure

Glutamate transporters constitute a distinct family of Na+-dependent transporters, with no significant homology to any other known protein family. Members are known from all three superkingdoms, Eukaryota, Bacteria and Archaea, with those found among the eukaryotes being specific for either glutamate/aspartate or neutral amino acid substrates. The EAATs (those specific for glutamate/aspartate) are present in multiple forms in many organisms, with humans, for example, containing five different paralogues fl'om this protein family. The five human EAATs share 40-60% amino acid sequence identity among themselves, whereas comparisons of the same type of transporter among different species of mammals yield much higher identities o{" approximately 90%. The molecular biology and pharmacology of EAATs in the mammalian CNS have been reviewed extensively (Takahashi el ell., 1997; Palacin el al., 1998: Vandenberg, 1998; Masson el ell., 1999; Seal and Amara, 1999: Slotboom et al., 1999; Danbolt, 2001; Gadea and gopez-Colome, 2001a). The global structure of the mammalian EAATs has also been found by analysis of peptide hydrophobicity to be highly conserved. The profiles suggest a common tertiary structure comprising eight transmembrane c~-helical domains (TMDs) with several fi-pleated pore-loop structures in the vicinity of cTAT Lt

60 b- 50 4{)

Frequenc> {kHz) FIG.

16 The mean response characteristics

of high-frequency T-neurons in

Scapteriscus borellii (Mason et al., 1998, fig. 6A). Reprinted with permission from the

Company of Biologists.

SOUND SIGNALLING IN ORTHOPTERA

205

that multiple units carried the audio frequency information, but that only one or at most a few units carried the ascending ultrasonic responses. Longer latency responses were detected to ultrasound, but these were likely to be in the descending fibres and related to the ultrasound startle response (Hoy el al., 1989). In S. abbreviatus, which is flightless, the omega neurons had similar tuning and response properties to those observed in the low-frequency omega neurons in S. horellii. One example of a high-frequency T-cell was recorded and it was homologous with the high-frequency T-cell in S. abbreviatus, but with thresholds of greater than 80dB SPL over the whole range of frequencies tested. The presence of responses to high-frequency in the night-flying S. borellii but not in the flightless S. abbreviatus, suggests that the auditory system in S. horellii plays a part in the avoidance of predators (see section 7.3). The two pairs of omega neurons found in the mole crickets are probably homologous with those of crickets, where two pairs are also found (Wohlers and Huber, 1982). Two pairs of omega neurons are also found in haglids (Mason and Schildberger, 1993), in contrast to the tettigoniids where only one pair has been found in the species studied so far (R6mer el al., 1988; Schul, 1997).

5.2.5

Audilo W interneurons in ,grasshoppers

The auditory pathways in the nervous system of grasshoppers have been studied extensively. There have been a number of reviews of auditory processing that include sections on the auditory interneurons, notably those of Stumpner and yon Helversen (2001), while the auditory processing of information from chordotonal organs is reviewed by Field and Matheson (1998). Excitatory and inhibitory inputs to two ascending interneurons were identified by Marquart (1985). Subsequently, Boyan (1991, 1992} showed that there was a common synaptic input from one of the cells recorded by Marquart onto other ascending auditory interneurons. One of these post-synaptic cells was the G neuron, also known as neuron 714, and this neuron is perhaps one of the best-characterized neurons in insects. A recent study of this morphologically prominent neuron by Boyan (1999) has demonstrated that the level of excitability of the neuron depends upon the gating of modulating inputs, including auditory and vibratory sources. Eight auditory interneurons are known to be presynaptic to neuron 714 and all are excitatory. However, it is not yet known how many of the receptors in the ears themselves connect synaptically with neuron 714. Those that do ascend to the metathoracic ganglion to synapse with neuron 714 seem to be predominantly high frequency (15 kHz) ones (Halex et al., 1988). The connections between identified cells in the metathoracic ganglion and neuron 714 in the mesothoracic ganglion are

206

D.J. ROBINSON AND M. J. HALL

summarized by Boyan (1999) and shown in Fig. 17. Neuron 714 is one of several interneurons that provide multimodal outputs in the form of combined information from more than one sensory source, and it is part of the neuronal chain that triggers jumping in locusts (Gynther and Peerson, 1989). The extensive range of both inputs and outputs to this neuron highlight its importance as a component of the locust nervous system and it is clearly a neuron which will repay deeper study in the future.

5.3 SYMMETRYANDASYMMETRY The sound-producing apparatus in Orthoptera shows asymmetry both in anatomy and in motion (for examples see Ewing, 1989). However, orthopteran ears would be expected to be symmetrical as directional information would be derived from the differences between the signals from the left and right ears. However, Rheinlaender and R6mer (1980) found asymmetry in the ears of the bushcricket Tettigonia viridissima when they investigated the responses of the T-cells, particularly at frequencies below 12kHz and above 20kHz. Of the

mesothoracicganglion

I

J,

.... t "£1:ta;h;;aci:;;gl~;n """ "~~

"" I'1" 1"] I

aft ~ " "

\-@ --@

~, @----....--~@1

,@ ,@,

,(

FIG. 17 A summary of the synaptic connections to neuron 714 in the locust mesothoracic ganglion, from cells in the metathoracic ganglion. The numbers in the circles refer either to identified interneurons or to the likely number of interneurons that make up the pathway, based on the measurement of synaptic delays. Arrows indicate an excitatory connection, filled circles an inhibitory one. Auditory afferents make a monosynaptic connection with neuron 714. Reprinted with permission from Boyan, G. S. (1999). Presynaptic contributions to response shape in an auditory shape in an auditory neuron of the grasshopper. J. Comp. Physiol. A 184, 279-294, Springer-Verlag GmbH & Co. KG.

SOUND SIGNALLING IN ORTHOPTERA

207

pairs of T-cells whose thresholds they measured, 53% showed some degree of asymmetry, explained in their view by a loss of function in single primary afferents. Boyan (1979) found an even larger proportion of individuals showing asymmetry of spiking responses in neurons in his study of Teleog~Tllus commodus. These results are puzzling when considering the importance of directionality, particularly in phonotaxis, and a recent study (Faure and Hoy, 2000a) has revisited the problem of the degree of asymmetry in the Tcell response, using Neoconocephalus e,siger. Of 32 males tested with a loudspeaker at 0 is strongly basic and should therefore form a strong ion pair. The active factor (F2, slower eluting) required four additional steps for isolation to homogeneity. Drying down samples in the presence of bovine serum albumin (BSA) to remove acetonitrile from the mobile phase was found, using RIA to quantify recovery, to lead to appreciable losses of sample. Esch el a/. (1983) advocate never drying down samples of bioactive neuropeptides during RPLC, but instead diluting fractions three to four-fold with water to decrease the acetonitrile concentration so that the sample will absorb to the column in the next RPLC step. Modification of the instrument to allow this approach was not possible with the instrument used, so a vacuum centrifuge was used to partially remove acetonitrile between purifications. However, this still led to rather poor recovery of F2, so that no estimates were made for the quantity of F1 and F2 per locust head (Schooley el al., 1987). Amino acid analysis of the two peptides gave identical amino acid compositions, suggesting they were homologues of AVP containing Leu and lie, but no Tyr or Phe. Each sample was reduced and carboxymethylated (RCM) prior to sequence analysis. Purification of the RCM peptides showed them to have identical retention on RPLC: sequence analysis of each showed that both peptides had the same primary sequence: C L I T N C P R G . This peptide was

296

G. M. COAST, I. ORCHARD, J. E. PHILLIPS AND D. A. SCHOOLEY

synthesized in both the C-terminal amidated and acid forms, which were reduced and carboxymethylated. The RCM-amide was found to co-elute with RCM-F1, while the RCM acid form had different retention properties, establishing the sequence as CLITNCPRG-NH2. Some of the remaining native F1 and F2 were analysed by size exclusion LC with a calibrated column; the amounts of peptide injected were too small to detect by UV, but were easily detectable in numerous timed fractions by RIA. This analysis showed F2 to have about twice the M,- of F1. Consequently, difficult specific syntheses of F2 were conducted as both parallel and antiparallel dimers. These were analysed by RPLC and the antiparallel dimer (the faster eluting of the two) was found to co-elute with F2. Bioassays using semi-isolated locust MTs including the ampullae and a portion of gut indicated that only the antiparallel dimer had biological activity. This established the 'AVP-like insect diuretic hormone' to be an antiparallel dimer of primary sequence C L I T N C P R G - N H 2 (Proux el al., 1987).

4.2.2

CRF-related neuropeptides

4.2.2.1 Isolation and puro?cation. M. sexta DH (Manse-DH) was isolated from 420 g of trimmed pharate adult heads (10 000 animals). The isolation was monitored with a slow, difficult assay in which aliquots of purification fractions were injected into decapitated, newly eclosed Pieris rapae butterflies (Kataoka et al., 1989). This species undergoes a post-eclosion diuresis (see section 3.3), which is blocked by ligaturing the neck of pharate adults, but is restored on injection of material with diuretic activity. A full 5nmol of peptide was obtained from 10000 animals and was easily sequenced, corresponding to about 0.5 pmol/head. The purification scheme utilised four RPLC purification steps and one ion-exchange LC step. The 41-residue peptide sequence has a 40% identity to sauvagine, a frog skin peptide, which at that time was thought to be the amphibian homologue of CRF. Interestingly, synthesis of the Cterminal free acid form of Manse-DH (as opposed to the amidated form) gave a product whose biological potency was reduced 1000-fold compared with Manse-DH in the P. rapae (and other) bioassays (see section 5.5.2). In 1991, Blackburn et al. isolated a second DH from M. sexta, which had escaped detection by Kataoka et al. (1989), using an in vivo bioassay with adult M. sexta. This peptide has only 30 residues and seems to represent a paralogue of Manse-DH. When aligned as shown in Fig. 4, only seven residues are identical between these peptides. The only major difference in biological potency between these two DH reported to date is that Manse-DH can promote fluid reabsorption from the everted rectal sac of M. sexta larvae, whereas ManseDPII cannot (Audsley et al., 1995). A further 11 CRF-related DHs have been isolated and sequenced (see Table 2 and Fig. 4), and an additional member of this family is encoded in the D. melanogaster genome (see below).

INSECT DIURETIC AND ANTIDIURETIC HORMONES

297 +

+ Musdo

DP

Pe~ am

gP

-

goone

DH+

-

Dippu

DH+

Locmi

DH Achdo D P Te m o D H ~ Manse

-

T G

S G

P

+ T G

A V

P

-

T G

. . . . -

XelJla C R F sauvacjlne

-

DPiI DH,

-

S

N KIP

T G P - M G MG - • G A Q AGALGESGA

DH

~; ...... D.

-

-

H v• ] l i DH: 1 Catco U I i, is an octapeptide with blocked N- and C-termini (see Table 4), which TABLE 4 The sequence of Manse-CAPeb compared with that of putative CAPehlike peptides from D. melanogaster and with identified PVKs. Bold type shows sequence identity to Manse-CAPxb Species M. sexm D. melanogaster P. [llll~'FiCdlla L. nladerae

L. migratoria

Peptide M anse-CAPeb Drome-CAP:~-I Drome-CAPeb-2 Peram-PVK- 1 Perarn-PVK-2 Leuma-PVK-1 Leuma-PVK-2 Leuma-PVK-3 Locmi-PVK- 1

Sequence

Reference

pELYA g PRVa GANMGLYAFPRVa ASGLVA F PRVa GASGL I PVMRNa GSSSGL 1S M PRVa GSSGL1 P FGRTa GSSGL I S MPRVa GSSGM I PFPRVa AAGL FQ F PRVa

Huesmann et al. (1995) Vanden Broeck (2001) Vanden Broeck (2001) Predel et al. (1995) Predel et al. (1998) Predel et al. (2000) Predel et al. (2000) Predel et al. (2000) Predel and Giide (2002)

308

G.M. COAST, I. ORCHARD, J. E. PHILLIPS AND D. A. SCHOOLEY

in addition to its cardioacceleratory activity, stimulates secretion by fruit-fly MTs (Davies el al., 1995), although it has no effect on hawkmoth tubules (N. Tublitz, personal communication}. Using in silico cloning, Vanden Broeck (2001) identified the D. melanogasler CAP2b gene (CG15520; capa), which localises to 99C8-99D1 on chromosome 3R. The gene encodes two putative CAP2b-like sequences (Table 4) with the same C-terminal as Manse-CAP2b (AFPRV), each followed by an amidation signal. The 151-residue prepropeptide begins with a 16 amino acid signal sequence and, in addition to the two CAP2b sequences, contains a 15-residue peptide flanked by cleavage sites (Lys Arg) and a C-terminal amidation signal. The C-terminus of this putative peptide (FGPRL-NH2) is characteristic of pyrokinin/pheromone biosynthesis activating neuropeptides (PBAN), which are known to have myotropic activity on cockroach hindgut (Holman et al., 1986b). It will be interesting to know whether this peptide has any effect on MT secretion. Manse-CAPzh-like peptides have also been identified in extracts of perivisceral organs from P. americana (Predel el al., 1998), k. maderae (Predel et al., 2000) and the migratory locust, L. mig, ratoria (Predel and Gfide, 2002). Unfortunately, these CAP2~-like peptides were christened periviscerokinins (PVKs), which, although reflecting their tissue of origin does not recognise their sequence similarity to Manse-CAP2b (see Table 4). The C-termini of Peram-PVK-1 and Leuma-PVK-1 are dissimilar from Manse-CAP2b, although their N-termini are strikingly similar to the CAP2b-like peptides of both species (see Table 4). 4.2.6

Tenebrio molitor and Leptinotarsa decemlineata A D F

Antidiuretic factors that inhibit MT secretion have been characterised from two coleopteran species. Lavigne el al. (2001} reported the purification of a factor from L. decemlineata through five RPLC steps using only two different columns; the protocol used involved minimal changes of parameters between successive purifications, and utilised evaporation steps between RPLC purifications likely to result in poor recovery (see section 4.2.1). While no peak was visible in the final chromatogram, they reported its apparent molecular size (estimated by dialysis) as 25-50 amino acids. Antidiuretic factors have been isolated and identified recently from pupal heads of T. molitor (the same stage of T. molitor used for isolation of Tenmo-DH3v and -DH47); the hydrophobicity of these factors (as reflected by retention behaviour on RPLC colmnns) and their size (13 and 14 amino acid residues) suggests they may be related to the L. decemlineata factor. The two ADF from T. molitor were isolated based on their ability to elevate cyclic GMP production by MTs (assayed using a commercial enzyme immunoessay (EIA)), rather than using a fluid secretion assay as did Lavigne et al. (2001). Fluid secretion assays for these ADF are difficult, because one


309

must quantify the decrease in basal secretion. The extraction procedures developed allowed a highly selective pre-purification from 1500 head equivalents. After fat removal with dichloromethane, heads were extracted with 90% aqueous methanol, which recovers a lot of protein but little activity. However, extraction of the pellet with a p H 4 acetate buffer gave good recovery of biological activity and relatively few impurities. This extract was pre-purified on a weak cation exchange cartridge (Toyopearl CM650M); the activity was retained upon loading the column with the 20 raM, pH 4 acetate buffer, but eluted on changing to a 2 0 m M , pH 7 acetate buffer. This highly enriched eluate required only three steps of RPLC on narrow-bore columns to obtain each factor in a pure state. The factors were sequenced, synthesised, and assayed for their effects on T. molitor tubules in secretion assays (laboratory of Dr S. W. Nicolson), The larger peptide, T e n m o - A D F a , has the sequence V V N T P G H A V S Y H V Y - O H and a calculated pl of 6.89 (Compute p I / M W tool at www.ExPASy.ch: the only ionic sites are two His residues and the free N- and C-termini). It has an exceptionally potent effect on T. molitor tubules (ECs0 10fM), but shows receptor desensitisation or internalisation at high (l nM and above) concentrations (Eigenheer et al., 2002). The smaller peptide, T e n m o - A D F b , has the sequence Y D D G S Y K P H I Y G F - O H and an ECs0 of 240pM in a fluid secretion assay, 24000-fold less potent than its congener. T e n m o - A D F b is also more acidic than its congener. (The former has a calculated pl of 5.17, which is more consistent with the poor solubility in acidic extracting solvent.) BLAST searches of these two sequences revealed that T e n m o - A D F a has an interesting similarity to the endothelins: 57% identity to rabbit big endothelin I, or 64% allowing for the conservative substitution of Val for Leu (see Fig. 6). However, it seems inappropriate to term T e n m o - A D F a an endothelin-like peptide, because the similarity is entirely to that part of endothelin which is removed in the proteolytic processing of big endothelin I to the potent vasoconstrictor endothelin 1. In fact, the N-terminus

. . . . 3880 7enmo J~J:)Fa

- A G Y H A P L V H[~Y A Y S A P[~F R A A T L S T V ~ A [ ~ I s [ ~ H V y . . . . . . . . . . . . . . . . . . . . . . . . . . VlV N T P G HIAIVlSIYIH V Y . . . .

Homsa ETZ Tenmo ADFb TmPCPg.2

. . . . . .

.

AGP

.

.

.

C S C SLSSJL M D K E CLVJY F C H L D I I ~,lV N T P]ELHJVlVlPJY GJL G S P - R S .

.

.

VAYAAVP

.

.

.

.

.

.

.

.

AGSGLEGQWI

.

.

.

.

.

PD

.

I NEKL

GI F

Y D O G S Y K P H

I IY

YDDGSYKPH

IYL~F

-

-

-

FIG. 6 Sequence alignment of T. molitor (Tenmo) ADFa and b, T. molitor cuticle proteins CAA03880 (40 C-terminal residues only) and TmPCP9.2 (40 C-terminal residues only), and rabbit (Oryctolagus cuniculus, Orycu-ET1) and human (HomsaETI) big endothelin I, aligned using Clustal W. Identical residues are boxed. Big endothelin I is cleaved by endothelin converting enzyme at the W V bond into the vasoconstrictor endothelin 1; the similarity to Tenmo-ADFa occurs in the inactive part of big endothelin I, on the C-terminal side of the cleavage site. The similarities were determined using the BEAST algorithm against the non-redundant database, but setting the expect value from 10 to 10000, which is important R~r short sequences.

310


of the cleaved, inactive piece of endothelin I is at the amino terminal residue of ADFa. Interestingly, Tenmo-ADFa is identical, except at residue 4, to the 14 C-terminal residues of a T. molitor cuticle protein (CAA03880) and TemnoA D F b is completely identical to the 13 C-terminal residues of T. molitor putative cuticle protein 9.2 (TmPCP 9.2; Baernholdt and Andersen, 1998; see Fig. 6). Because of the far lower potency of Tenmo-ADFb, and its disturbing complete identity to a cuticle protein of this species, its structure was not submitted for publication until immunocytochemical evidence was available that it appears to be produced mainly in two pairs of bilaterally symmetrical cells in the protocerebrum (Eigenheer et al., 2002b). This evidence is consistent with a bonaJide neuropeptide role for Tenmo-ADFb, although its relative lack of potency compared with Temno-ADFa may mean its primary role is something other than antidiuresis. About 33 fmol of A D F a was recovered per head (Eigenheer et al., 2002a), compared with c.200 fmol Tenmo-ADFb (Eigenheer et al., 2003). In contrast, the same pupal heads of T. molitor contain c.45 fmol per head of Tenmo-DH37, although head extracts have only a diuretic effect (Wiehart et al,, 2002). This is again consistent with the desensitisation phenomenon observed in the response to the ADF, in contrast to relatively low desensitisation with CRF-like DH. 4.2.7

M. sexta antidiureticJactors

Liao et al. (2000) demonstrated the presence of antidiuretic activity in brain CC-CA complexes of M. sexta that stimulates fluid uptake from everted rectal sac preparations of larval M. sexta. Fractionation of an aqueous extract of 300 neuroendocrine complexes on a polymeric RPLC column gave two zones of activity that stimulated fluid reabsorption. The more abundant, slower eluting of these was christened Manse-ADFB. This factor appears to have much in common with the action of Schgr-ITP on the ileum of S. gregaria in that it promotes active CI- transport (see section 5.4). Interestingly, basal reabsorption from everted rectal sacs is blocked by addition of bafilomycin A~ or by amiloride, drugs which block MT secretion (see section 2.2), but these have no affect on the ADFB-stimulated fluid reabsorption (see section 5.4). This blockage of basal reabsorption allows improvement of the everted rectal sac assay for isolation; inclusion of amiloride in the bathing medium makes the effect of ADFB more evident so smaller doses can be used. Manse-ADFB was purified to partial homogeneity from an 80% methanol extract of 10 000 frozen larval heads of M. sexta; the choice of solvent was based on poor recovery of activity from tissues extracted with acidic solvents. The extract was concentrated to remove methanol, defatted, and pre-purified by absorption to a ToyoPearl QAE-550C anion-exchange cartridge. The 0.25 M NaC1, pH 8, eluent was separated by two successive semi-preparative RPLC steps, then by an anion-exchange LC step, followed by three additional RPLC steps. The apparently homogeneous product had an M,- of 8770.6 Da, in


311

the same range as the neuroparsins but c.200 Da higher than Schgr-ITP. Only partial sequence data were obtained (Liao, 2000). Based on this a cDNA clone from a brain CC CA c D N A library was isolated and sequenced: the peptide encoded by this clone was unfortunately a M. sexta homologue of cytochrome bc~ subunit H, rather than an ADF. The deduced protein sequence matched the partial protein sequence, showing that the putative ADF sequenced was an impurity, probably masking a far lower quantity of bioactive peptide (S. Nagata and D. A. Schooley, unpublished data). No attempts have been made to isolate the less abundant ADFA, and neither was it characterised to the degree that ADFB was. 4.2.8

Mosquito natriuretic peptide ( M N P )

Three fractions were isolated from a head extract of A. aegypti by RPLC on the basis of their effect on the TEP of isolated perfused MTs (Petzel et al., 1985). Fraction I depolarised the TEP, but had no effect on fluid secretion, although it increases tritiated water (THO) loss and urine output from intact flies, possibly by inhibiting fluid uptake from the hindgut (Wheelock et al., 1988). Fraction I! also depolarised the TEP, whereas the response to fraction Ill was biphasic, with the TEP first depolarising and then hyperpolarising. Fractions II and Ill both have diuretic activity and selectively stimulate secretion of NaCl-rich urine. Fraction Ili was the more potent, and its diuretic and natriuretic activity was indistinguishable from that of exogenous cyclic AMP, although the latter only hyperpolarised the TEP (Petzel et al., 1985). All three fractions contain peptides with Mr estimated by gel-filtration chromatography of 2425 (fraction 1), 2721 (fraction II) and 1862Da (fraction III) (Petzel et al., 1986). Fraction II1 was named Mosquito Natriuretic Peptide (MNP) and was shown subsequently to stimulate cyclic AMP production in isolated tubules (Petzel et al., 1987). 4.2.9

F. polyctena antidiureticjitctor ( F o p A D F )

The MTs of forest ants (F. polyctena) recently collected from the nest fi'equently fail to secrete and have a closed lumen (Van Kerkhove et al., 1989). One day later, tubules from the same batch of ants have an open lumen and are secretory, leading to the suggestion that an A D F is released in response to stress (Laenen et al., 2001). Using a 15% trifluoroacetic acid (TFA) extract of 150 000 ant abdomens, gaenen et al. (2001) isolated an ADF that reversibly inhibits MT secretion by 70%. Biological activity was lost towards the end of the purification by RPEC, but the presence of an ADF in a 15% TFA extract of haemolymph from ants with non-secreting MTs suggests it has an hormonal function. The active factor was christened FopADF, and Laenen et al. (2001) suggest that it is released when the poison gland is emptied. Since this gland contains an estimated 10% of the total body

312


water, it may be necessary to inhibit MT secretion to conserve water reserves, which are then replenished by drinking. 4.3

PURIFICATION AND CHEMICAL STRUCTURE OF NEUROPEPTIDES T H A T STIMULATE LOCUST H I N D G U T

4.3.1

Introduction

Early studies identified the retrocerebral complex, consisting of the median neurosecretory cells (MNCs) of the pars intercerebralis (PI) and their axons projecting to storage and release sites in the CC, as the major source of stimulants of locust hindgut. However, initiation of stimulant release from the CC using K + depolarisation in the presence of Ca 2+ has not been successful to date (J. E. Phillips and J. Meredith, unpublished observations). Stimulatory activity is also present in ventral ganglia VG4 to VG7 of locusts (Lechleitner and Phillips, 1989; Audsley and Phillips, 1990). The locust VG stimulant has different properties from that of the CC with regard to the time course for changes in the hindgut Isc, solubility, acid lability and heat stability. Bilgen (1994) partially purified the acid-labile VG factor, which had an approximate mass by size-exclusion chromatography of 37 000 Da, i.e. several times that of factors from the CC discussed below. Three neuropeptides have been purified fully or partially from the CC of locusts (reviewed by Phillips and Audsley, 1995; Phillips et al., 1998a,b): Neuroparsins, Ion Transport Peptide (ITP), and Chloride Transport Stimulating Hormone (CTSH). Only the first two have been fully sequenced. Neuroparsins and CTSH were bioassayed on recta, whereas ITP was bioassayed originally on ilea. 4.3.2

Neuroparsins

Herault et al. (1985) reported that the CC and the glandular lobe (GCC) of the L. migratoria CC both contain a rectal ADH factor, each differing in size and extraction properties. The GCC factor was not purified, but Herault and Proux (1987) report that GCC extracts cause a sharp peak in rectal tissue cyclic AMP levels coinciding with elevated J,.. This stimulation is mimicked by forskolin, a stimulant of adenylate cyclase. The factor from the CC was identified as neuroparsins, because all antidiuretic activity in crude CC of L. migratoria was abolished by an antibody to this neuropeptide (Fournier and Girardie, 1988). Neuroparsins are two proteins (NpA, NpB) isolated and sequenced from CC of L. migratoria by Girardie et al. (1989, 1990). NpB is a homodimer of a 78-residue polypeptide (8188 Da). NpA is identical to NpB except for an additional heterogeneous Nterminus, the longest of which has 83 residues. NpB is thought to be formed from NpA by cleavage of the terminal amino acids. However, Hietter et al.


313

(1991) have proposed a revised monomeric structure for neuroparsins involving three disulphide bridges. Fournier (1991) has reviewed pharmacological and direct evidence that NpB acts on rectal J,. by stimulating the inositol phosphate (IP) cascade, resulting in elevation of cystolic Ca 2+. In summary, they conclude that neuroparsins are the only A D H in L. migratoria CC and that these peptides act via the 1P Ca 2+ second messenger system rather than via the cyclic AMP pathway. They did not study actions of purified or synthetic Nps on rectal solute transport processes. However, rectal Jv in the absence of (or indeed against) an osmotic difference across this epithelium must of necessity be driven secondarily by solute transport. Since stimulation of rectal J, is abolished in CI--free saline, antidiuretic factors such as Nps presumably must first act by stimulating the predominant ion transport process, CI- absorption. Neuroparsins might possibly also increase osmotic and cation permeability to enhance fluid transport. Some comment on the rectal Jv bioassay used by Fournier and Girardie (1988) is in order, because their results appear to differ from those reported later. These workers pre-incubate everted rectal sacs in CI -free saline for 1 h (Fournier el al., 1987), a treatment known to cause drastic loss of most CI from this tissue (Williams et al., 1977). They then restore preparations to normal C1 -containing salines, with or without stimulants or other test agents on the haemocoel side. This ionic change in itself stimulates considerable increase in Jv of the controls over the next hour. The effect of stimulants was assessed from a small additional increase in Jv in the same 1 h time period. Since the major ion pump in locust rectum is an apical C1 pump, restoring CIto C1 -depleted cells introduces a transient situation in which increased C1entry (with K +) into rectal cells with accompanying fluid would be expected to cause cell swelling. This in turn could trigger well-known cell volume regulatory mechanisms in which elevated cell Ca 2+ initiates KC1 and hence fluid exit from these cells. Thus it is not clear whether neuroparsins stimulate long-term steady-state J,. (i.e. as measured over several hours when C1- is always present) or some short-term and transient (1 h) cell volume regulatory response. Subsequently, Jell; and Phillips (1996; see also Jeffs, 1993: Phillips et al., 1998b) observed no effect of Nps over several hours on either rectal J,., or on rectal and ileal Isc of the desert locust even at high doses. An unlikely explanation might be the presence of different major stimulants in the CC of these two locust species. However, reciprocal bioassays of CC extracts from L. migratoria and S. gregaria on rectal and ileal 1so and J, in these two locust species have been more recently conducted (Macins et al., 1999). Corpora cardiaca extracts from either species were equally effective in several hindgut bioassays on both species of locust, suggesting similar stimulants are present. L. migratoria CC must therefore contain a major stimulant of S. gregaria rectal lsc and Jv other than Nps because the latter had no action on S, gregaria hindgut (Jeffs, 1993; Jeffs and Phillips, 1996). Moreover, the deduced amino acid sequences of the

314


ITP neuropeptide from the CC of S. gregaria and L. migratoria are identical (Macins et al., 1999), while their ITP-like (ITP-L) neuropeptides differ only by one neutral amino acid substitution (see section 4.3.4.6). A neuroparsin cDNA from L. migratoria was recently expressed using a baculovirus vector (Girardie et al., 2001) and the peptide found to have a small stimulatory effect on rectal J,, using the questionable bioassay discussed above. Further studies of neuroparsin actions on other well-characterised transport bioassays are required to clarify the conflicting reports on its stimulatory role on hindgut. 4.3.3

Chloride Transport Stimulating Hormone ( C T S H )

The CI -dependent l~c across flat sheet preparations of S. gregar& recta has been used as a bioassay to partially purify a neuropeptide stimulant from the CC (Spring and Phillips, 1980a,b). This preparation maintains a 10-fold increase in Isc for >8 h after stimulation with cyclic AMP or CC extracts. An active factor (CTSH) eluted as a single peak with an apparent Mr of about 8000Da using a size-exclusion (BioGel P-30) column (Phillips et al., 1980; Phillips el al., 1982; Phillips et al., 1986). Biological activity is destroyed by trypsin digestion, indicating that the active factor is a peptide. The estimated concentration of CTSH required to cause maximum increase in rectal Is~ is ,

K

t

r, >,:,e ~:P : :,,~ ;t v,r E~; ,r,J }-

K+ Na*

NH4*v/o

midline~ l

Segmental boundary

,

,~ r

B

C

FIG. 23 Locmi-DH-like immunoreactivity in the abdominal segment of L. miL,ratoria. A The Locmi-DH-like immunoreactive posterior lateral cells of thc abdominal ganglia project out through the sternal nerve, resulting in neurohaemal areas of the perivisceral organ (PVO) of the transverse nerve, paramedial nerve and lateral heart nerve. Redrawn from Patel et al. (1994). B Whole mount preparation of abdominal ganglion, illustrating the immunoreactive posterior lateral cells (PLC) projecting through the sternal nerve (SN). Note the immunoreactive projections centrally to the neuropile. C Immunoreactive projections along the oviducal nerve (OVN) which branches from the sternal nerve of abdominal ganglion 7. Photomicrographs kindly provided by Rodney Kwok. The scale bar shown in (B) represents: A, 350/~m: B, 50Hm; C, 10#m.


355

nerves, and at the junction of the transverse and paramedial nerve, with processes running along the nerves. In addition, immunoreactivity is also found in processes from the VIIth abdominal ganglion which project over the lateral oviducts (Fig. 23). In a similar fashion, CRF-like immunoreactivity is found in the MNCs and CC of M. sexta and R. prolixus (Veenstra and Hagedorn, 1991; Emery et al., 1994; Te Brugge et al., 1999). In R. prolixus, the immunoreactive processes also extend through the CC to the aorta (Te Brugge et al., 1999). Within M. chmTestica (laboni et al., 1998), the MNCs are also Locmi-DH-like immunoreactive, although in the larva they lie in a lateral position in the brain. Projections from these NSCs pass to the ventral sector (equivalent to the CC) of the ring gland (a complex structure incorporating the CC, CA and prothoracic gland) in the larva and to the CC in the adult. Also in M. s e x m and R. prolixus (Fig. 24) there is a prominent group of intensely immunoreactive posterior lateral NSCs associated with the abdominal ganglia or neuromeres. In M. sexta larvae, Manse-DPII-like immunoreactivity is found in these posterior-lateral NSCs with their axons projecting out of the ventral nerve into the next transverse nerve, where they branch to produce neurohaemal terminals (Emery et al., 1994). A prominent pair of neurons reactive against Manse-DH in the posterior portion of the terminal abdominal ganglion have axons which project out of the ipsilateral eighth ventral nerve to innervate the rectum and provide neurohaemal-like terminals in the cryptonephric complex (Chen et al., 1994b). In starved larvae, a pair of ventral, paramedial cells are also Manse-DH-like immunoreactive. These cells are believed to be the so-called M4 NSCs, which project anteriorly through the median procurrent nerve to the transverse nerve. These M4 cells also stain positively in adult M. se.vta, as do the posterior lateral NSCs with projections to the transverse nerve via the ventral nerves, where they produce neurohaemal-type terminals (Chen et al., 1994b; Emery et al., 1994). Within R. prolixus (Te Brugge et al., 1999), 10 12 posterior-lateral NSCs of the M T G M project through abdominal nerves 1 and 2 and produce positively stained neurohaemal areas lying on the surface of these nerves (Fig. 24). The immunoreactive processes can also be traced to the body wall, where staining is evident around the spiracles. Fine immunoreactive axons are also seen in abdominal nerves 3 5 and in the genital nerve. The situation in adult M. domestica is somewhat different, in that there is a fused thoracic abdominal ganglion with one pair of Locmi-DH-like immunoreactive cells located ventrally in each neuromere (laboni et al., 1998). lmmunoreactive processes from these NSCs project to the dorsal neural sheath overlying the entire ganglionic mass, where they form an extensive plexus of varicose Locmi-DH-like immunoreactive terminals. Midgut endocrine cells in R. prolixus stain weakly for Locmi-DH-like immunoreactivity (Te Brugge et al., 1999), although midgut endocrine cells, as well as endocrine cells in the ampullae of L. migratoria, stain positively for

356


A

MTGM It,

P L C / f " ~',':.:'~:~:

J

\NH

FIG. 24 Whole mount preparations of leucokinin-like and Locmi-DH-like immunoreactivity in two species of bugs, R. prolixus and Oncopellux fasciatus. A and D Leucokinin-like immunoreactivity in the mesothoracic ganglionic mass (MTGM) of R. prolixus. Note the immunoreactive posterior lateral neurosecretory cells (PLC) with central projections as well as projections out through abdominal nerves. Neurohaemal (NH) areas appear on the surface of abdominal nerves. The PLC are also immunoreactive for Locmi-DH-like immunoreactivity (not shown). B and C Higher magnification of posterior lateral neurosecretory cells (PLC) within the equivalent ganglion of O../ilscialus stained for leucokinin-like immunoreactivity (B), and axons from other lateral neurosecretory cells of O. jasciatus which possess apparent reservoirs of Locmi-DH-like immunoreactivity prior to their exit along abdominal nerves (C). These cells in O../asciants do not co-localise Locusfa-DH-like and leucokinin-like immunoreactivity. Photomicrographs kindly provided by Victoria Te Brugge. Scale bars: A. 100#m: B and C, 25#m; D, 15/zm.


357

CRF-like peptides in A. aegypti (Veenstra et al., 1995) and L. migratoria (Montuenga et al., 1996). With regard to hindgut, Locmi-DH-like immunoreactive processes are present over the entire structure in R. prolixus (Te Brugge el al., 1999). At the ultrastructural level, Locmi-DH-like immunoreactivity is associated with dense-core neurosccretory granules (diameter 450 nm) of the endocrine cells in the ampullae of L. migratoria (Montuenga et al., 1996), and in a variety of morphologically distinct electron-dense granules in R. prolixus (Te Brugge et al., 1999). In R. prolixus, immunogold electron microscopy has been performed on the CC, aorta and abdominal nerves and reveals Locmi-DH-like immunoreactive material overlying the electron-dense neurosecretory granules located within the terminals (Te Brugge et al., 1999). In the CC, two types of granules are evident: round granules of approximately 73nm diameter: and oval granules, 121 x 74nm. In the aorta, only oval granules are immunoreactive, whereas the abdominal nerves again contain round granules approximately 110 nm in diameter. The presence of morphologically distinct immunoreactive granule types suggests the presence of multiple members of the CRF-related family. Structural evidence for this is available from other insects (see section 4.2.2.2) where different structural forms of the CRF-like diuretic peptides are present within an individual species. The distribution of CRF-like immunoreactivity (see above) and the presence of Locmi-DH in the PI and CC of locusts, as revealed by MALDIT O F MS (Matrix-Assisted Laser Desorption Ionisation Time-of-Flight Mass Spectrometer) (Clynen et al., 2001), is consistent with this family of peptides having a neurohormonal function in insects. In addition, Locmi-DH is released by high-K + saline (Audsley et al., 1997b: Clynen et al., 2001) from the CC of L. mi~,ratoria in a Ca:+-dependent manner (Audsley et al., 1997b). Moreover, Locmi-DH can be detected in the haemolymph of fed locusts (Patel et al., 1995), and anti-Locmi-DH antiserum specifically blocks Locmi-DH-induced diuretic activity in vivo, thereby preventing an increase in primary urine production in recently fed locusts (Patel et al., 1995: see section 7.1.3).

6.5

KINJNS

Detailed maps of the distribution of kinin-related peptides within the nervous system of a variety of insect species have been developed using antisera generated against insect kinins, most notably Leuma-K-l. Kinin-like immunoreactivity is distributed extensively throughout the nervous system of a wide range of insect species, and within a range of neuron types, including interneurons, NSCs and possibly sensory cells. Interestingly, although the distribution of kinin-like immunoreactive neurons in abdominal ganglia is very similar in all species examined, there are differences in the type and number of immunoreactive neurons in the brains of different species (see

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G . M . COAST, I. ORCHARD, J. E. PHILLIPS AND D. A. SCHOOLEY

N/issel, 1996c). Thus, there is apparently no leucokinin-like immunoreactivity in the brains of A. mell(/'era (Chen et al., 1994a); very few immunoreactive neurons in mosquitoes or flies such as D. melanogaster and Calliphora vomitoria (N~issel, 1993a; Kim, 1998), but large numbers in Phormia terraenova, L. m~ratoria, L. maderae, Spodoplera ]ittorina and R. prolixus (see Nfissel, 1993a; Te Brugge et aL, 2001). Consistent throughout the species examined is the presence of kinin-like immunoreactivity in NSCs and their concomitant neurohaemal areas. Interestingly, within this consistency, two variables have been noted by Chert et al. (1994a). The first is the degree to which the MNCs of the PI of the brain are kinin-like immunoreactive, and the second variable is the location and the extent of immunoreactive neurohaemal organs associated with the NSCs of the abdominal ganglia (see Chen et al., 1994a). This latter variability has been noted earlier for abdominal neurohaemal organs in general (see Grillot, 1983). In L. maderae, approximately 100 MNCs are found to stain intensely for leucokinin-like immunoreactivity in the PI of the protocerebrum (N'assel et al., 1992). These MNCs send axons to the CC via the paired NCC 1. In addition, there are approximately seven lateral neurosecretory cells (LNCs, dorsolateral in the protocerebrum) that are positive for leucokinin-like immunoreactivity, and which project axons via the NCC 2 to the CC. The MNCs and LNCs of L. maderae supply the storage lobes of the CC with large amounts of immunoreactive neurohaemal terminals, lmmunoreactive axons also continue on into the hypocerebral ganglion and then through the oesophageal nerve and recurrent nerve into the neuropil of the frontal ganglion. The immunoreactive axons then continue through the two frontal connectives into the tritocerebrum, where they produce swollen neuropilar terminals. Leucokinin-like immunoreactive MNCs and LNCs are also found in the PI of the cockroach Nauphoeta cinerea, the cricket A. domesticus and the mosquito, A. aegypti (Chen et al., 1994a). In contrast to the situation described above, an anti-Leuma-K-IV antiserum usually does not stain the MNCs of either larval or adult brains of M. sexta, but occasionally weak staining is observed in the larval group lla MNCs and adult IIb MNCs, with projections passing into the CC (Chen et al., 1994b). Within the CC, however, the intrinsic NSCs are intensely immunoreactive, as they are in the cockroach N. cinerea (Chert et al., 1994a,b). In a similar manner, the MNCs of R. prolixus typically do not stain for leucokinin-like immunoreactivity, but in some preparations from insects that have been starved for 10 weeks, there is staining in a subset of MNCs (Te Brugge et al., 2001). Under the conditions tested, there is no leucokinin-like immunoreactivity in the MNCs or NCC 1 of L. migratoria (N/issel, 1993a,b), S. americana, A. melllT"era (Chen el al., 1994a) or M. domestica (Iaboni el al., 1998). Another dipteran, however, P. terraenovcte, does express leucokinin-like immunoreactivity in the MNCs (sce N~ssel and Lundquist, 1991).


359

As mentioned earlier, the distribution of leucokinin-like immunoreactive neurons in abdominal ganglia is very similar in all species examined, although the distribution of their neurohaemal areas can differ. In all insects studied so far, sets of posterior lateral NSCs of the abdominal ganglia are leucokinin-like immunoreactive (see reviews by Chen et al., 1994a; Nfissel, 1996c). Typically, the abdominal ganglia 1 7 contain a set of leucokinin-like immunoreactive posterior lateral NSCs with axons projecting out through the posterior nerve. Again, typically, the last 1, 2 or 3 abdominal neuromeres lack these positively stained posterior lateral NSCs. The number of posterior lateral NSCs per ganglion varies between insects (from 1 to 7 pairs) and can also vary between ganglia of the same insect. In L. maderae, there are two pairs of leucokinin-like immunoreactive NSCs in each unfused abdominal ganglion, and, in R. prolixus, 6-7 pairs in the abdominal neuromeres of the M T G M . Similar bilaterally paired clusters of varying numbers of cells have been shown for N, cinerea (Chen el al., 1994a), P. americana (Agricota and Brfiunig, 1995), L. m&,raloria (Thompson el al., 1995), S. americana (Chen el al., 1994a), A. domesticus (Chcn et al., 1994a), Gryllus himaculatus (Helle el al., 1995), Agrotis segetum (Cantera eta/., 1992), M. se.vta (Chen et al., 1994a), A. mell(fera (Chen et al., 1994a), A. aeg)7~ti and Phalacrocera replicata (Cantera and N/_issel, 1992: Chen el al., 1994a), D. melanogaster, C, vomitoria, P. terraenovae, M. ~kmwslica (Cantera and Nfisscl, 1992; Iaboni el al., 1998) and R. prolixus (Te Brugge et a[., 2001). Processes from the leucokinin-like immunoreactive posterior-lateral NSC extend out of the posterior nerve in L. maderae and continue into the link nerve and then the dorsal segmental nerve of the next segment (N~ssel et al., 1992). The axons then project to the lateral cardiac nerve on the ipsilateral side. Prior to entering the dorsal segmental nerve, some axons branch and produce varicose processes to the main tracheal trunks near the spiracles. Also, there is a branch from each axon from the link nerve, which passes to the transverse nerve of that segment, thereby reaching the pcrisympathetic neurohaemal organ. Immunoreactive varicosities and terminals are found in the processes near the spiracles, in the lateral cardiac nerve, and in perisympathetic organs, indicative of neurohaemal release sites. A similar peripheral distribution of lcucokinin-like immunoreactive processes and neurohaemal sites has been described for N. citwrea (Chen et al., 1994aL L. mi~,ratoria (Thompson eta/.. 1995), A. domesticus (Chen el al., 1994a) and G. himaculatus (Helle el al., 1995). with additional neurohaemal sites in the sheath of the root of the dorsal nerve of the next posterior segment in the latter two species. Within larval 4. segettml, the abdominal posterior lateral NSCs send axons to the alary muscles, spiracles and transverse nerves; and, in the adult, the processes in the transverse nerve project to the neural sheath on the dorsal surface of the abdominal nerve cord and form an extensive immunorcactive plcxus (Cantera et al., 1992). The projections from abdominal NSCs in larwd and adult M. se.vta pass out of thc posterior nerve and form varicose processes in the neurohaemal region of the transverse nerve of the next posterior segment

360


(Chen et al., 1994a). The processes are more intensely stained and greater in number in the adult. In A. mell(/'era, the neurohaemal sites are located in the segmental nerve root near the ganglion, whereas in larval D. melanogaster, C. vomiloria and P. terraenovae the leucokinin-like immunoreactive processes supply terminals to the abdominal body wall muscles: and in the adult, they produce neurohaemal sites on the pericardial septum (Cantera and Nfissel, 1992), Within the M. domestica adult, there are seven to ten pairs of ventrolateral leucokinin-like immunoreactive neurons and a pair of dorsomedial neurons in the abdominal neuromeres of the thoracico-abdominal ganglionic mass (Iaboni et al., 1998). Seven or eight of the ventrolateral neurons have axons which project out of the two fused lateral abdominal nerves, where they produce immunoreactive neurohaemal sites. The dorsomedial neurons project out the median abdominal nerve and again result in immunoreactive varicosities on the nerve (laboni et al., 1998). In A. aegypti, the release sites appear to be on the surface of the segmental nerves (Chen et al., 1994a) and in A. mell(/i~ra there is a bulbous neurohaemal organ on the surface of the ventral nerve root near the ganglion (Chen et al., 1994a). Abdominal nerves 1 and 1I possess leucokinin-like immunoreactive neurohaemal sites originating from the posterior-lateral NSCs of the mesothoracic ganglionic mass in R. prolixus (Te Brugge eta/., 2001). Projections from these neurons also extend to the lateral margin of the abdomen, producing a complex branching pattern associated with the tergal sternal muscles (Te Brugge et al., 2001). In addition, there are paired multipolar peripheral neurons immunoreactive for leucokinins on branches of peripheral nerves in R. prolirus. The presence of kinins in apparently homologous posterior-lateral NSCs of abdominal ganglia of a wide variety of insect species has been suggested to be a highly conserved feature in insects (Chen et al., 1994a). The distribution of the abdominal neurohaemal areas associated with such cells reveals some variation, although the extensive neurohaemal areas produced are consistent with kinins being released into the haemolymph as neurohormones, Despite the presence of leucokinin-like immunoreactivity in NSCs and neurohaemal organs throughout the insect species studied (and by inference their potential as neurohormones), it is still worth exploring the possibility of innervation and release over target organs. Initial studies in L. maderae found no such innervation to the MTs, hindgut or fat body (N/issel et al., 1992), but leucokinin-like immunoreactive processes do project to the foregut (crop) of this species, and immunoreactive bipolar and tripolar neurons are also present in the crop. Using a different antiserum, one generated against Leuma-K-VIII, Meola et al. (1994) found projections of leucokinin-like immunoreactive processes from the posterior nerve of the terminal abdominal ganglion, which might indeed project to the hindgut. In R. prolixus, no kinin-like immunoreactive processes are seen projecting from the stomatogastric nervous system to the foregut, although the midgut does contain leucokinin-like immunoreactive endocrine cells, and leucokinin-like immunoreactive processes produce


361

a plexus of staining over the posterior midgut and hindgut (Te Brugge et al., 2001). Similarly, in A. aegypti and the crane fly, P. replicata, paired leucokininlike immunoreactive neurons in the terminal ganglion project to the hindgut (Cantera and N'assel, 1992). Within the terminal abdominal ganglion of M. s e x t a larvae, there are two leucokinin-like immunoreactive neurons (ventral unpaired median (VUM) neurons) with kinin-like immunoreactive axons projecting out of the terminal nerves, innervating the posterior hindgut and also forming an apparent neurohaemal area in the cryptonephric complex (Chert et al., 1994b). These two VUM neurons could not be distinguished in the adult. lmmunoreactive staining of NSCs and neurohaemal areas is certainly consistent with the antigen being a neurohormone, released from these structures into the haemolymph. This is further corroborated by the demonstration by HPLC and RIA or M A L D I - T O F of the presence of the kinins within neurohaemal areas and their release from such structures. Thus, all eight isoforms of the leucokinins have been shown to be associated with the CC and lateral heart nerves in L. maderae (Winther et al., 1996), and various P. americana kinins are associated with the different neurohaemal areas of that cockroach (Predel el al., 2001). In addition, in A. domesticus and L. maderae, kinin-like immunoreactivity has been found in the haemolymph, and in both these species Ca 2 ~dependent release of kinin-like material from the CC has been demonstrated in response to depolarisation with high K + saline (Muren et al., 1993: Chung et al., 1994: see section 7.1.4).

6.6

CAP2b/PERIVISCEROKININS

As mentioned earlier (see section 4.2.5), Manse-CAP2b, was originally sequenced from M . s e x t a (Huesmann el al., 1995) and belongs to a group of at least five peptides (CAPs) that collectively influence cardiac function and patterns of behaviour in M . se_vta (Tublitz et al., 1991). These peptides may be derived from midline NSCs that project to the abdominal transverse nerve perisympathetic organs (see Wegener et al., 2001). Manse-CAP2b belongs to a family of peptides which have been isolated and sequenced from cockroaches and locusts, and found within the D. m e h m o g a s t e r genome, and which share the common C-terminus PRVamide (see Table 4; section 4.2.5). It is of some interest that a sequence comparison of the cockroach PVKs (see Table 4: Wegener el al., 2002) illustrates similar N-terminal sequences, but variable C-terminal sequences. This is unusual for neuropeptide families, which tend to share the C-terminal sequence that typically contains the active core. Thus, while the PVKs (and Manse-CAP2b) that share the PRVamide C-terminus are biologically active on MTs, there is no evidence that other members of the PVK family are also active. For example, 0.1/IM Peram-PVK-1 has no effect on A. domesticus tubules, whereas 0.03#M Peram-PVK-2 has significant

362

G . M . COAST, I. ORCHARD, J. E. PHILLIPS A N D D. A. SCHOOLEY

diuretic activity (G. M. Coast, unpublished observations). Since the immunohistochemical staining of insect nervous systems has used antibodies generated against the different PVKs, it is difficult to generalise on the distribution of those PVKs that may act as diuretic/antidiuretic peptides. With that caveat in mind, however, PVKs have been shown to be present in the anterior median nerve and abdominal perisympathetic organs of P. americana using MALDITOF MS (Predel, 2001). The ratio of peptides present was 2:1:0.2 for PeramPVK-I, Peram-PVK-2 and Leuma-PVK-2 respectively, although MALDI-MS is notoriously non-quantifiable. In contrast, the posterior median nerves do not have any detectable PVKs, and neither do the thoracic perisympathetic organs, nor the retrocerebral complex. Using antisera specifically generated against either Peram-PVK-1 or Peram-PVK-2, Eckert el al. (1999) and Predel el al. (1998) found these peptides to coexist in immunoreactive clusters of 6 10 NSCs in abdominal ganglia. In the five unfused abdominal ganglia, the cell bodies occur as three clusters (Ci C3) and as two cells ventrolateral to the C~ cluster. Axons from these cells project through the anterior median nerve to the perisympathetic neurohaemal organs, where they produce intensely stained neurohaemal areas. Processes continue on to innervate the hyperneural muscle, and, via the link nerve to the segmental nerve, project to the heart, alary muscles and segmental vessel (Eckert et al.. 1999). The fused terminal ganglion reflects a similar pattern of immunoreactive cells in only the seventh neuromere (Predel et al., 1998; Eckert el al., 1999). The more extensive study, using Peram-PVK-1 antisera (Eckert el al., 1999), reveals an intrinsic neuronal network of immunoreactive cell bodies and processes within the brain, SOG and metathoracic ganglion, and local interneurons in the proto- and tritocerebrum, again indicating that these neuropeptides play a central role, as well as acting as neurohormones. It should be remembered, however, that Peram-PVK-I is not known to have activity on MTs. A peptide with similar chromatographic properties and biological activity to M. sexta CAP2~, has been isolated from D. melanogaster (Davies et al., 1995), and two related peptides are predicted from the fruit-fly genome (Vanden Broeck, 2001; Wegener et al., 2002). CAP2h-like bioactivity in D. melanogaster has been found in a set of midline mesodermal cells that have axonal-like processes in the transverse nerve, suggesting a secretory function for the cells (see Huesmann et al., 1995). The PVKs, as defined by Wegener et al. (2002) are among the most abundant peptides stored in the abdominal perisympathetic organs of P. americana, as revealed by M A L D I - T O F MS (Wegener et al., 1999; see Wegener el al., 2001). Approximately 6 pmol Pea PVK-I is stored in these organs in P. americana. Similarly, the distribution and abundance of Manse-CAPeb in M. s e x t a abdominal perisympathetic organs is comparable with that of the cockroach (Wegener et al., 2001). Evidence for release of PVKs from these organs comes from K + depolarisation experiments. Thus, approximately 28 50fmol Peram-PVK-1 is releasable in a Ca2+-dependent manner, fl-om an abdominal nerve cord with intact perisympathetic organs


363

that has been exposed to a saline containing 100 mM K + (Wegener el al., 2001). To date, however, PVKs have not been shown to be present in the haemolymph. 6.7

( ' A L C I T O N I N - L I K E PEPTIDES

Antisera to the insect calcitonin-related diuretic peptides have only recently become available, and there are no published reports on their distribution. Preliminary results (V. A. Te Brugge, personal communication), using antisera against Dippu-DH31 (Furuya el al., 2000b; produced by V. C. Lombardi and D. A. Schooley), indicate neurohaemal sites on the abdominal nerve in D. punctata. Within R. prolixus, Dippu-DH31-1ike immunoreactivity is found throughout the CNS, with fine processes observed in some of the nerves of the M T G M . Weak staining of neurohaemal sites is also observed on abdominal nerve 2. Future work is needed to create a detailed map of the distribution throughout a variety of insect species. 6.~

ION TRANSPORT F'EFq-IDE

Neurosecretory cells immunoreactive to an antiserum raised against Orconectes limosus C H H originate in frontal mediolateral brain areas of L. migratoria and have branches within the superior median protocerebrum. Their axons run through the NCC I to the CC and then to the CA in the nervi corporis allati I (H. Dircksen, personal communication). The discrete localisation suggests this antisermn recognises ITP rather than ITP-L, and ITP m R N A is known to be expressed in the brain and CC (Meredith el al., 1996). Similar results (M. Fuse, personal communication) were obtained with an Schgr-ITP antiserum, which detected small numbers of immunoreactive NSCs in the P1 and CC of S. gregaria. As with the Orcli-CHH antiserum, it is not clear whether the ITP antiserum also recognised ITP-L. Additionally, there are three to ten C H H immunoreactive NSCs in each thoracic and abdominal PVO of L. migratoria and sometimes also in the link nerves (H. Dircksen, personal communication). Meredith et al. (1996) did not detect ITP m R N A in the VG of S. gregaria, but the preparation that they used may not have included the PVOs. The presence of C H H (ITP)-like immunoreactive material in the CC is consistent with it being released into the circulation although, to date, release of ITP from the CC has not been demonstrated and it has not been definitively identified in hacmolymph. 6.9

( O-LOCALISATION

The posterior lateral NSC which stain for kinin-like immunoreactivity within the abdominal ganglia of a variety of insects, have an identical cell body position and morphology to those that stain for CRF-like D H immunoreactivity.

364


Double-label immunohistochemistry confirms that in L. migratoria (Thompson et al., 1995), M. sexta (Chen et al., 1994b) and R. prolixus (Te Brugge el al., 2001), kinin-related and CRF-related diuretic peptides are co-localised to the same posterior lateral NSCs in the abdominal ganglia and their neurohaemal endings (in M. sexta only one (L3) in each of the two pairs of (L3 4) of leucokinin-like immunoreactive cells co-localise Manse-DH). This discovery led Thompson et al. (1995) to suggest that co-localisation within the posterior lateral NSCs of abdominal ganglia may be a conserved feature among insects. However, laboni et al. (1998) could not detect co-localisation of these two peptide families in either brain or abdominal neuromeres in M. domeslica. Moreover, despite co-localisation of Locmi-DH-like and Leuma-Kl-like peptides in the posterio>lateral NSC in the blood-feeding bug, R. prolixus (Fig. 24), co-localisation is not evident in the milkweed bug, Oncopehus Jitsciatus (V. A. Te Brugge, personal communication). Thus, as commented on by Te Brugge el al. (2001), co-localisation of CRF-related and kinin-like diuretic peptides in posterior lateral NSC of abdominal ganglia may well be a 'general phenomenon in insects' (Thompson el al.. 1995), but not necessarily a universal one. It is unclear if this co-localisation extends to the MNCs in the brain. Fewer insects have been examined for CRF-related DH immunoreactivity, and kininlike immunoreactivity has only been shown consistently in the MNCs of L. maderae, N. cbwrea, A. domesticus and A. aegypti. Unfortunately, corresponding double labelling experiments with the CRF-related diuretic peptide antisera have not been reported for these species. Within R. prolixus and M. sexta, although the MNCs in the brain stain intensely for the CRF-like DH immunoreactivity, the staining for Leuma-K-like immunoreactivity is seen in only a few preparations, which makes double-labelling immunohistochemistry hard to interpret (see Chen et al., 1994b: Te Brugge et al., 2001). In those R. prolixus preparations where there are Leuma-K-I-like immunopositive medial NSCs, these cells are single labelled for Leuma-K-I-like immunoreactivity (Te Brugge et al., 2001). However, other cell types (approximately 24-28 cells in the brain, two to four in the prothoracic ganglion, 12 18 in the M T G M ) are double-labelled for Leuma-K-l-like and Locmi-DH immunoreactivity (Te Brugge et al., 2001). Within L. mi,~ratoria, the posterio~lateral NSCs of abdominal ganglia which stain for Leuma-K-l-like immunoreactivity label with antisera to Locmi-DH-like immunoreactivity (see above), but also with lysine vasopressin (LVP, Thompson et al., 1995). Similar results are found for posterior lateral abdominal NSCs in L. maderae, N. cinerea and A. domesticus (Niissel el a[., 1992). N'assel et al. (1992) consider that the co-localisation with LVP-like immunorcactivity in L. maderae is due to cross-reaction bctwcen the LVP antiserum and the endogenous leucokinins, since preabsorption of the LVP antiserum with Leuma-K-1-BSA conjugate abolishes tissue staining. However, staining in L. migratoria is not blocked by such preabsorption


365

(Thompson et al., 1995), and these authors conclude, along with additional evidence from H P L C and immunoassay, that the abdominal ganglia in L. mi~raloria contain three distinct epitopes reacting with the three antisera. The medial NSCs in the PI of L. migratoria which are Locmi-DH-like immunoreactive are also immunoreactive to antisera generated against ovary maturing parsin, a putative neurohormonc which stimulates oogenesis in locusts (Tamarelle e l al., 2000). Other forms of co-localisation have been shown in P. americamt, where Peram-PVK-1, Peram-PVK-2 and Peram-pyrokinin-5 appear to coexist in the abdominal nerve cord and interneurons of the brain (see Wegener el al., 2001). The expression of these peptides starts at the same time in embryonic development, leading Wegener el al. (2001) to suggest that both PVKs and Peram-pyrokinin 5 are encoded together on a single gene in P. americmza as they have been shown to be in D. melam)¢asler (Vanden Broeck, 2001: see section 4.2.5). The M4 NSC in M. sexta appears to co-localise Manse-DHlike, PBAN-like and CCAP-Iike immunoreactivity (Chen el al., 1994b). Also, the SOG cells in L. migraloria which stain for Locmi-DH-like immunoreactivity, and which appear to be the same cells that have previously been shown to project to the heart, would be expected to co-localisc with FMRFamide-like immunoreactivity (Patel el al., 1994). Of considerable interest is the study by Zitnan el a/. (1995), which reports a map of NSCs in the brain of M. sexta that stain for prothoraciotropic hormone, bombyxin, allatotropin, allatostatin, DH, eclosion hormone and proctolin. Such a map allows inferences to be drawn about possible co-localisation with other neuropeptides when performing immunohistochemical studies in this species. Although there is no evidence for the co-localisation of diuretic peptides with serotonin within NSCs or neurohaemal terminals, these two groups of diuretic factors are co-localised in a number of interneurons within the various ganglia of R. prolixus (Te Brugge et al., 2001 ). It also seems reasonable to conclude that the wtrious diuretic factors might be expected to be co-localised within interneurons of other insects as well. To sum up, a common, although not universal feature in insects (see laboni et al., 1998; Te Brugge et al., 2001) is the co-localisation of kinin-like and C R F like diuretic peptides in the abdominal ganglia posterior lateral NSCs and their neurohaemal terminals. Such co-localisation does not seem to occur in the M N C s in the PI of insects studied, although a broader range of species needs to be examined. Other peptide phenotypes, for which there is no evidence of diuretic activity, may also be co-localised with the diuretic peptides in NSCs. Serotonin does not appear to be co-localised within NSCs that express diuretic peptides. It would appear that there is sufficient flexibility in NSC types for the kinins, CRF-like diuretic peptides and serotonin to be released independently into the haemolymph, or, at least in the case of kinins and CRF-like diuretic peptides, to be co-released from the same NSCs. In addition, there is the possibility that

366


these factors can be co-released or released independently within the neuropil of the central ganglia where they might modulate the activities of central neurons, which might well be involved in behaviours other than diuresis. The presence of immunoreactivity in processes lying over the digestive tract certainly implies physiological activities for these diuretic factors, which goes beyond diuresis itself, and suggests a broader role for them in feeding behaviour as a whole. It is interesting to note the antifeedant activities reported for Manse-DH in H. virescens larvae (Keeley et al., 1992), Manse-DPll in M. sexta neonates (Ma et al., 2000) and Locmi-DH in fifth-instar L. mi~gratoria nymphs (Coast and Goldsworthy, 1997; see section 9.2).

7

Physiological relevance

7.1

7.1.1

C I R C U L A T I N G LEVELS IN R E L A T I O N TO P H Y S I O L O G I C A L STATUS

hTtroduction

The identification of factors with diuretic or antidiuretic activity using in vitro bioassays is but a first step towards showing that they have a functional role as circulating neurohormones. Importantly, it is necessary to show that they are released from neurohaemal sites into the circulation and that the haemolymph concentrations achieved are appropriate for mediating a physiological response. This necessitates measuring haemolymph concentrations and investigating the effects of injecting putative hormones into intact insects (see section 7.2.3). Unfortunately, little information is available in either of these areas and, to date, there is no evidence to show that ITP, the only well characterised stimulant of hindgut reabsorption, has a hormonal function. 7.1.2

Serotonin

Feeding induces release of serotonin from an extensive network of serotoninlike immunoreactive neurohaemal areas and processes that arise primarily from the M T G M and supply the dorsal and ventral integument (Orchard et al., 1988; Lange el al., 1989). Within minutes of the onset of feeding in fifthinstars, serotonin is released from these abdominal nerve processes both at the integument, probably participating in cuticular plasticisation (Reynolds, 1974, Orchard et al., 1988), and into the haemolymph, resulting in a 15-fold increase in serotonin concentration (Lange et ell., 1989). The titre of serotonin within the haemolymph of fifth-instar R. prolixus is shown in Fig. 25. As can be seen, the concentration of serotonin in the haemolymph of unfed insects is low (6.8 nM), but rapidly increases following the onset of feeding, peaking to 115 nM within 5 min. Thereafter the titre declines, and remains low, but still elevated over unfed controls, for 24h. Adult male R. prolixus also have


367

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Time after c o m m e n c e m e n t of feeding (min) FIG. 25 The titre of serotonin in the haemolymph of fifth instar R. proli.vu.~ at varying times after the commencement of Feeding on rabbit blood. Time zero represents unfed insects. Symbols represent means ±S.E. Redrawn from Lange ~'f al. (1989).

elevated serotonin levels within 3 min of the onset of feeding, with a peak concentration somewhat less than in fifth-instars, but with the same pattern of sustained elevation and decline towards baseline levels (Barrett et al., 1993). The serotonin that appears in the haemolymph is not derived from the blood meal, since similar titres are found when fifth-instars are fed on an artificial diet. Rather, as referred to above, the serotonin appears to be released from the peripheral nervous system, since there is a depletion of the serotonin-like immunoreactive staining of abdominal neurohaemal areas and processes over the body wall during feeding (Lange el al., 1988; Orchard el ol., 1988), and a depletion in content of serotonin as determined by HPLC with electrochemical detection. In addition, serotonin is released from these neurohaemal areas (Lange el al., 1988) in response to a depolarising stimulus (high K" saline). The peak titre of serotonin in the haemolymph of R. prolixus is sufficient to induce cyclic A M P elevation and fluid secretion from the MTs it7 vilro (Maddrell et al., 1971; Barrett ~,t al., 1993), and anterior midgut (crop)

368


(Farmer et al., 1981; Barrett et al., 1993), leading to the conclusion that serotonin is a neurohormone in R. prolixus, released by the natural stimulus of feeding, and controlling diuresis. It has been proposed that serotonin has a broad range of peripheral roles that result in it being a co-ordinator of feedingrelated activities in R. prolixus, including plasticisation, salivation and gut motility, as well as fluid secretion from crop and MTs (see Orchard et al., 1988; Barrett and Orchard, 1990; Barrett et al., 1993; Orchard and Brugge, 2002). 7.1.3

A VP-like insect D H

Changes in the haemolymph titre of AVP-IDH in adult L. migratoria over 24 h are shown in Fig. 26, along with an index of primary urine production based upon measurements of amaranth clearance (Picquot and Proux, 1987). The concentration of AVP-IDH fluctuates between 0.04 and 0.4 nM and, in insects that were fed daily, there is a peak at 12:00 h, which coincides with a peak in primary urine production. At other times, however, the two parameters vary independently of one another, which argues against any causal relationship. In contrast to the data of Picquot and Proux (1987), Baines et al. (1995) found very little (c.2 pM) AVP-like material in the circulation. Arguably, AVPIDH may be neither a diuretic nor a neurohormone: it has no effect on secretion by isolated tubules (see section 5.2.2); the haemolymph titre is low for a peptide hormone; and it is debatable whether there are neurohaemal sites for its release (see section 6.1.3). 7.1.4

C R F - r e l a t e d peptides

Using a highly sensitive RIA, Audsley et al. (1997b) showed that Locmi-DH is mainly stored in the brain and CC, with small amounts (a few 100 fmol) in thoracic and abdominal ganglia. In addition, a small amount of Locmi-DH is present in endocrine cells of the MT ampullae (Montuenga et al., 1996). Feeding causes an immediate release of Locmi-DH from the CC, which can be mimicked by depolarising the glands in high K + saline (Audsley et al., 1997b). The haemolymph titre of Locmi-DH increases throughout the meal (see Fig. 27: Audsley et al., 1997a), which normally lasts about 15 20rain. During this time, the amount of peptide stored in the CC returns to pre-feeding levels, reflecting its continued export from the brain, and Locmi-DH synthesis by MNC may be upregulated (Audsley et al., 1997b). Locmi-DH immunoreactive axons follow a spiral course within the brain (see Fig. 22) and contribute to the 'neuropilar reservoir' described by Highnam and West (1971). This reservoir empties of neurosecretory material within 5 rain of the onset of feeding (Highnam and West, 1971), which possibly reflects the export of Locmi-DH to the CC.


6-

369

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40% of the excess salt and water are excreted within 1 2 h (Williams et al., 1983). In the peak phase of diuresis, urine output can reach 8 0 n L m i n -1 (see Fig. 3) and diuretic activity is detectable in the haemolymph by bioassay (Wheelock et al., 1988). Significantly, the cyclic AMP content of the MTs is increased 10-fold (Petzel et al., 1987), which is consistent with the release of a diuretic peptide (MNP?) that activates this second messenger pathway. During the peak phase of diuresis, drops of NaCl-rich urine isosmotic to haemolymph are voided every 30s (see Fig. 3) and little or no modification occurs in the hindgut (Williams el al., 1983). Thereafter, urine output declines dramatically, but remains higher than in unfed insects, which void little or no urine. At the same time, the urine changes from being NaCl-rich to KCl-rich as excess K + from the imbibed blood cells is voided (Williams et al., 1983). Importantly, the urine is now hypo-osmotic to mammalian plasma, probably


379

due to NaCI uptake from the hindgut, which allows osmotically free water to be cleared and prevents dilution of the haemolymph. The switch from natriuresis to kaliuresis could be due in part to the disappearance of M N P from the circulation and the release of kinins, which do not have natriuretic activity (Pannabecker et al., 1993). Unlike mosquito tubules, which secrete NaCI-rich urine in the peak phase of diuresis, significant amounts of K + are secreted from serotonin-stimulated distal tubules of R. prolixus (see section 5.2.1.2), sufficient to deplete the haemolymph of K + within a minute at high urine flow rates (see Maddrell et ul., 1993)! This is prevented by K - reabsorption from the proximal tubule (Maddrell and Phillips, 1975), which is activated prior to distal tubule secretion (see Maddrell el ul., 1993). Since both the proximal and the distal tubule are maximally stimulated throughout diuresis, K + homeostasis depends upon their a u t o n o m o u s response to changes m haemolymph [K +] (Maddrell e! al., 1993). This is shown for A. domesticus tubules in Fig. 28. As can be seen, any change in bathing fluid [K +] is countered by a change in K + secretion, especially in Achdo-DP-stimulated tubules, and this is likely to be important for excreting excess K - after a meal. In the two to three hours after a blood meal, a volume of fluid equal to five to ten times the haemolymph volume is absorbed from the midgut of R. prolixus and voided via the excretory system (see Maddrell, 1980). To preserve

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[K*]bf (mM) FIG. 28 The effect of bathing fluid [K ~] on K + transport by A. domestictts tubules before (dotted line) and alter (solid line) stimulation with 50 nM Achdo-DP. Symbols represent means ±S.E. Cricket haemolymph contains 7.6 mM K ~ (shown by the vertical line), and any departure from this value is automatically countered by an increase or decrease in K ~ transport, most notably in the presence of Achdo-DP. G. M. Coast, unpublished data.

380


the haemolymph volume, it is necessary to balance fluid movement across the midgut and MTs, and Maddrell (1980) proposed that this be achieved with a DH that activates both processes, but with differing potency and efficacy. Assuming that the hormone has greater potency on MTs, but greater efficacy on the midgut, any imbalance is automatically rectified. For example, if MT secretion exceeds midgut absorption the haemolymph volume will fall, which will increase the concentration of D H in the circulation and further stimulate midgut transport. Serotonin might serve this purpose in that it stimulates fluid transport >six-fold across the isolated anterior midgut of fifth-instar nymphs (EC5o 50nM; Farmer el al., 1981), but is a more potent stimulant of distal tubule secretion (EC50 30~40nM; Maddrell el al., 1993). Serotonin has also been shown to stimulate ion transport by midgut and MTs of A. aegyt)ti larvae (Clark and Bradley, 1996; Clark el al., 1999), and may therefore be used to coordinate their activities when the insects are transferred to saline media (Clark and Bradley, 1997). Urine output declines dramatically from 4 0 0 n L m i n l to < 5 0 n L m i n t towards the end of the postprandial diuresis in R. prolixus (Maddrell, 1964a). Quinlan el al. (1997) suggest that release of a Manse-CAP2b-like peptide may contribute to this, since CAP2b has been shown to reduce secretion by serotonin-stimulated tubules by activating a cyclic GMP-dependent cyclic AMP phosphodiesterase (see section 5.2.5). In support of this, they show that the decline in urine output at c.3 h post-feeding correlates with a 56% increase in tubule cyclic G M P content. 8.3

SYNERGISM BETWEEN D I U R E T I C H O R M O N E S

The dose-response curve for Locmi-DH is shifted to the left and made steeper when tested in the presence of a low concentration of 0.05 nM Locmi-K (see Fig. 29; Coast, 1995), which is evidence of synergism. This is a predicted outcome of the peptides acting via different second messenger pathways to stimulate different transport pathways (see Clark el al., 1998a), and has also been demonstrated in housefly tubules with Musdo-DP and Musdo-K (Holman et al., 1999). Interestingly, there is no evidence of synergism between Achdo-DP and Achdo-K-I (Coast and Kay, 1994), which is consistent with the CRF-related peptide stimulating both cation and anion transport by A. domesticus tubules (see section 5.2.3.2). More surprising, however, Dippu-DH4{, and Dippu-DH31 act synergistically on D. punctata tubules (Furuya el al., 2000b), although they each use cyclic AMP as a second messenger and might therefore be expected to have an additive effect on tubule secretion. Similarly, the synergism between serotonin and either forskolin or a peptidergic D H from the M T G M in R. prolixus (Maddrell et al., 1993a) is unexpected, because both activate adenylate cyclase (Maddrell et al., 1993a). O'Donnell and Spring (2000) suggest that the synergism between diuretic factors that use cyclic AMP as a second messenger might result either from activation of different

INSECT DIURETICAND ANTIDIURETIC HORMONES 125-

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log c o n c e n t r a t i o n (nM) FIG. 29 Synergism between Locmi-DH and Locmi-K. The dose response curve for Locmi-DH (dotted line, triangles) is moved to the left and made steeper when tested in combination with 0.05 nM Locmi-K (solid line~ squares). Symbols represent means ±S.E. and the vertical lines show the haemolymph titre of Locmi-DH in fed and starved insects. Redrawn from Coast (1995).

isoforms of adenylate cyclase or through cross talk between second messenger pathways, and this clearly warrants further investigation. The peptidergic DH of R. pro/i_rus that synergises with serotonin has not been identified, but it is unlikely to be either a kinin or a CRF-related peptide although both are present in the M T G M (see sections 6.1.4 and 6.1.5). Kinins have no effect on secretion (Te Brugge eta/., 2002) and, in separate studies, neither Locmi-DH (Coast, 1996) nor Z o o n e - D H (Te Brugge eta/., 2002) has been shown to act synergistically with serotonin. There must therefore be some other peptidergic D H in the M T G M . Synergism produces an increase in potency (see Fig. 29), which means that less D H needs to be released. This permits diuresis to be switched off rapidly, because there is less D H to remove from the circulation. Using data from Coast (1995), it would require the release of about 90% of the Locmi-DH stored in the brain and CC (c.4nmol: Audsley et al., 1997b) to increase MT secretion from 10% to 90% of its maximum rate. The same effect can be achieved with the release of just 5% of stored peptide in the presence of 0.05nM Locmi-K. Synergism also makes the dose response curve steeper (see Fig. 29) allowing precise control of diuresis, because small changes in D H concentration have a large impact on tubule secretion. The independent and controllable release of the two hormones would further increase the precision with which tubule

382


secretion is regulated, and it is noteworthy that although kinins and CRFrelated peptides co-localise in abdominal ganglia NSCs, this has not been reported for MNCs of the brain (see section 6.1.9). 8.4

CO-ORDINATING MALP1GHIAN TUBULE AND HINDGUT ACTIVITIES

Fluid recycling between MTs, hindgut and haemolymph increases several fold in recently fed locusts (Phillips and Audsley, 1995), allowing toxic waste to be eliminated rapidly from the circulation while minimising excretory water loss. Malpighian tubule secretion and hindgut reabsorption must therefore be coordinated, which might be achieved with one hormone controlling both processes, as proposed for the integration of midgut and MT function (see section 8.2). Coast et al. (1999) conducted reciprocal in vitro bioassays with pure peptides to show that Schgr-ITP does not influence locust MT secretion, and stimulants of tubule secretion (Locmi-K and Locmi-DH) do not stimulate either rectal or ileal ls~. or J,.. Thus, hormonal control of these two major segments of the locust excretory system is clearly separated. This may be needed for the precise regulation of haemolymph volume and composition, but requires the release of DHs and ADHs to be co-ordinated. Unfortunately, nothing is known about the circulating levels of ADHs, although the amount of CTSH-Iike activity in haemolymph increases after feeding (Spring and Phillips, 1980c). As mentioned previously, excretory water loss is the difference between the rates of fluid entry into the hindgut and reabsorption in the ileum and rectum. The capacity of the locust ileum and rectum to reabsorb fluid could make it difficult for the insect to deal with an hydration stress without reducing fluid uptake in the hindgut, and one way that this might be achieved is with the release of an A D H antagonist, such as Schgr-ITP-L (Phillips et al., 1998b). The natural function of ITP-L is unknown, but the identical N-terminal sequences of ITP and ITP-L to residue 40 suggest that it might bind to the ITP-R and act as an antagonist of ITP so as to curtail hindgut fluid reabsorption, or act to reduce synthesis and release of ITP at the brain-CC level. Phillips et al. (1998b) suggest that excess reabsorption of dilute fluid in the hindgut due to overstimulation by ITP might cause general tissue swelling. This might directly trigger widespread release of ITP-L fi'om various tissues to reduce ITPstimulated fluid recovery in the hindgut leading to diuresis. In this sense, ITP-L might be analogous to vertebrate atrial natriuretic peptide (ANP), which is found in several body tissues and which inhibits renal salt and hence fluid reabsorption. This working hypothesis might now be examined using expressed ITP-L and antibodies available to this peptide. Sf9 cells transfected with baculovirus do not correctly cleave expressed ITP and ITP-L, leading to an 11 amino acid extension of the N-terminus, which could cause the antagonistic action of ITP-L on the ileal I~c bioassay. This possibility was excluded by Pfeifer el al. (1999) and Wang et al. (2000), who


383

used Drosophila Kcl cells and a designed plasmid vector to express ITP (KcITP) and ITP-L (KclTP-L) with correct N-terminal cleavage. Wang el al. (2000) found that 5 nM KclTP-L caused 75% inhibition of ileal I~c stimulation by 0.08 nM KclTP. Thus the physiological function of ITP-L proposed above remains a viable one for future study.

9

The excretory system as a target for pest control strategies

Hormones control many aspects of the physiology, and the behaviour of insects and the endocrine system is seen as an important target in the development of novel, safe and specific insecticides (Masler e t a / . , 1993: Kelly et al., 1994). Of the numerous endocrine targets available, the control of water balance is particularly attractive because the adults of all major pest species are terrestrial and must carefully regulate excretory water loss. For example, the use of DH agonists and/or A D H antagonists could lead potentially to dehydration and death. The excretory system is also an important route for the clearance of toxic waste, and if this is compromised, t"o1"example with a DH antagonist, it is likely to adversely affect survival. Maeda (1989) inserted a synthetic gene encoding Manse-DH into the genome of a baculovirus. The gene contained an N-terminal signal sequence based upon a D. melanogaster cuticle protein and a sequence that encoded Manse-DH extended by a glycine residue for C-terminal amidation. Northern blot analysis of m R N A from fat body of silkworm (Boml)y.v mori) larvae infected with the recombinant virus (BmDH5) confirmed the synthetic gene was expressed. Moreover, haemolymph collected from these larvae contained several hundred times more diuretic activity, as determined by the stimulation of post-eclosion diuresis in decapitated newly emerged P. rapae butterflies, than haemolymph from an equivalent number of M. se.vta pupae (Maeda, 1989). Critically, however, diuretic activity was not measured in haemolymph from B. moll larvae infected with wild-type virus, which might have confirmed the expression and secretion of correctly processed Manse-DH. A detailed analysis of large (c.50 mE) volumes of haemolymph from B. mori infected with BmDH5 showed the presence of a number of peaks on RPLC with diuretic activity, but none had a retention time coincident with ManseDH (C. A. Miller, R. G. Troeschler, S. J. Kramer, and D. A. Schooley, unpublished data). It seems likely that the biologically active zones observed in haemolymph from infected animals could well represent endogenous B. mori kinins, or other factors. Nevertheless, larvae infected with BmDH5 suffered a 30% reduction in haemolymph volume compared with controls and with larvae infected with wild-type virus (Maeda, 1989). Interestingly, water lost from the haemolymph of BmDH5 injected larwm is retained in the midgut, which is consistent with Manse-DH acting on both the cryptonephric and free portions of the MTs to recycle fluid from the rectal complex to the midgut (see

384


section 2.3). Significantly, all BmDH5 infected larvae were dead within four days, whereas >95% of larvae infected with the wild-type virus survived (Maeda, 1989). The cause of death was unclear, but it could have resulted from a build up in the concentration of toxic waste in the haemolymph as its volume declined (see Maeda, 1989). Ma et al. (2000) investigated the effect of administering Manse-DPI1 to M. sexta neonates by applying it to tobacco leaf discs at doses of between 1.5 ng and 15 000 ng. The larvae gained less weight, developed more slowly and suffered higher mortality than controls, although it is unclear what caused the latter. Surprisingly, Manse-DPII had no effect when injected direct into the haemocoel, which suggests it may have an antifeedant effect. Thus, Keeley et al. (1992) showed that Manse-DH has antifeedant activity in H. virescens larvae, and Locmi-DH has a similar effect in L. migratoria (Coast and Goldsworthy, 1997), but in both of these studies the peptides were injected into the haemocoel. At present, it is unclear whether Manse-DPIl applied to leaf discs is assimilated in a biologically active form, although this would be surprising given that it is likely to be broken down by proteolytic enzymes in the gut. Seinsche et al. (2000) has shown that injections of 50 pmol Helze-K-I into fifth-instar H. virescens larvae along with 1 tzmol captopril (an ACE inhibitor) results in 83% larval mortality within 5 days, compared with 7% in salineinjected controls and 44% in animals injected with the kinin alone. The scissile bonds of insect kinins (see section 7.3.4) can be protected with a sterically hindered anainoisobutyryl (Aib) residue. The Aib residue is compatible with a type VI fl-turn and FF-Aih-WG-NH2 retains the potency and activity of the parent compound (FFSWG-NH2) in a cricket tubule bioassay, but is resistant to ACE (Nachman et al., 1997). The double Aib analogue Aib-FS-Aib-WGNHz is completely resistant to degradation by H. zea tubules, and is more potent (ECs0 1.2pM) than native Achdo-Ks (ECsos of between 22 and 324pM) in the cricket assay (Nachman et al., 2002a). This analogue is also active in a housefly tubule assay (Nachman et al., 2002a), but is less potent (ECs0 1.5#M) than Musdo-K (ECs0 0.13 nM), consistent with the role of the N-terminal region for high affinity receptor binding (Coast et al., 2002; see section 5.5.3). Nevertheless, injections of 50pmol Aib-FS-Aib-WG-NH2 and Musdo-K are equally effective in stimulating radiolabelled inulin excretion from intact houseflies at 1 and 2h post-injection, most likely because the peptidase-resistant analogue survives longer in the circulation. While these results demonstrate the feasibility of developing stable kinin analogues, a key to their use as pest control agents is that they should be topically active, i.e. able to penetrate the waxy epicuticle. Teal and Nachman (1997) have developed amphiphilic analogues of PBAN that have pheromonotropic activity when applied topically in water to H. virescens cuticle. These analogues have a hydrophobic moiety replacing the Phe residue in the PBAN active core (FTPRL-NH2) to counter the polar side-chain of Arg. The hydrophobic


385

moieties tested included 6-phenylhexanoic acid, 9-fluorenacetic acid and 1-pyrenebutyric acid. The same moieties have been attached to the N-terminus of a kinin analogue, A R F F P W G - N H 2 , the Arg residue being included to produce an amphiphilic molecule. All three analogues retained full diuretic activity in a cricket tubule assay (ECs0s 0.25 nM to 1.7nM; R. J. Nachman and G. M. Coast, unpublished observations), but the lack of a reliable in viro assay has prevented them from being tested after topical application. In a search for novel agonists/antagonists at DH receptors, a tripeptide combinatorial library constructed from D-amino acids has been screened in a cricket tubule assay (G. M. Coast and R. J. Nachman, unpublished observations). Several agonists with the general sequence Fmoc-dPro-dPro-dXxx-NH2 were identified, all of which form a nascent right-handed polyproline lI helix that can be superimposed on the kinin type VI fi-turn conformation (Moyna el al., 1999). Since these analogues are constructed from D-amino acids, they are resistant to peptidase attack and could therefore be important leads for future studies. To summarise, CRF-related peptides and kinins have been shown to disrupt insect growth and development and increase mortality. This may not result from their diuretic activity, because increased secretion of primary urine is probably countered (automatically?) by fluid reabsorption in the hindgut. It is unclear whether effects on feeding behaviour are mediated peripherally or centrally, although the latter would require that they cross the blood brain barrier. Increased visceral muscle activity in response to CRF-related peptides (Blake et al., 1996) and kinins (Holman et al., 1990) might disrupt the passage of food through the gut and reduce feeding, while the antifeedant activity of Locmi-DH is attributed to an effect on gustatory sensilla (Coast and Goldsworthy, 1997). The identification of ITP receptor antagonists (see section 5.5.5) also offers a promising area for future research by inserting their cDNAs into insect viruses, and trial studies have already been conducted (D. Theihnann, Agriculture Canada; personal communication) using locust ITP cDNAs provided by H. W. Brock, J. Meredith and J. E. Phillips (University of British Columbia).

10

Future directions

While MT and hindgut epithelial transport processes are reasonably well understood in some insects, it is very evident from this review that the field of endocrine control of MT and of hindgut function in particular is still at an early stage. It is equally clear that a comprehensive understanding of the excretory system's role in whole body osmotic and ionic homeostasis will depend firstly on establishing which factors normally control MT secretion and hindgut reabsorption and secretion in ~,ivo and secondly on elucidating

386


how their release is controlled and co-ordinated. This is a challenge for the next generation. To date, five different neuropeptide families have been shown to have diuretic or antidiuretic activity on isolated MTs, but the peptidergic hormone of R. prolixus that acts synergistically with serotonin has not been identified, and MNP and FopADF have still to be sequenced. Similarly, a number of distinct neuropeptide stimulants of locust hindgut (e.g. CTSH, Ventral Ganglia Factor (VGF), a second stimulant of ileal l~c in HPLC factions from CC) and of the M. se.vta cryptonephric complex (Manse-ADF A and B) still have not been sequenced because of inadequate biochemical separation methods for small amounts of acid-labile peptides. For such peptides, expression-cloning and identification of secreted stimulants in functional bioassays appears to be the most promising approach to obtain deduced amino acid sequences from their cDNAs. Given the size of some of these neuropeptides, their production for physiological and structure activity studies, and for generating specific antibodies, is more easily done using a good expression system (e.g. in the Kcl-plasmid vector system and in protease-deficient S. cerevMae) when a cDNA has been obtained. Development of new bioassay systems for insects other than locusts and the tobacco hornworm will be essential, and this requires that the search for stimulant cDNAs should be directed to large insects within any new taxonomic group. An exception is D. melanogaster, where the completed genome, extensive stocks of genetic strains, and genetic manipulations offer unique potential (see Dow and Davies, 2001). The identification of genes encoding DHs and ADHs and their receptors permits functional studies (functional genomics) by observing the resultant phenotype after the gene of interest is mutated (reverse genetics) or "knocked out' by double-stranded RNA (dsRNA)-mediated interference. For example, Macins et al. (1999) found that Schgr-lTP-like activity appeared abruptly in the head, thorax and abdomen of embryos about 80% of the way through development in locust eggs. In order to study the functions of this neuropeptide during development, the Drome-ITP gene is being inactiwtted using transposons (H. W. Brock, J. Meredith and J. E. Phillips, unpublished observations). Such treatment is lethal. Conceivably Drome-ITP controlled hindgut fluid reabsorption causes the body volume expansion necessary for emergence of the first instar. Chung et al. (1999) have provided the first evidence that an ITP holnologue, CHH, has such a role during ecdysis in crustaceans. An alternative genome-wide approach is the use of DNA micro- or oligonucleotide arrays to monitor the expression profiles of all identified genes. Quantitative comparisons of expression profiles for genes encoding neuropeptides, their receptors, signalling pathways and ion transport/channel proteins in flies reared under different environmental conditions will provide a unique insight into the multidimensional control of excretory processes. The complementary technique of peptidomic analysis uses a combination of RPLC


387

and MS to identify peptides present in biological samples from their unique M,-s (Clynen et al., 2002). By comparing the haemolymph complement of neuropeptides in insects held under hydrating and dehydrating conditions it should be possible to identify, those components that change and might therefore have a physiological role in the control of salt and water balance. The Drosophila genome has been scanned for peptide GPCRs, and the majority, possibly all, have been identified (Hewes and Taghert, 2001). There appear to be about 20 orphan receptors. It should therefore be relatively easy to identify the receptors for Drome-DH31, Drome-CAP2b and Drome-lTP from binding of the radiolabelled ligands to these orphan receptors expressed in a suitable cell system. In turn, the deduced amino acid sequences for the receptors would permit production of antibodies against unique regions that could be used to identify potential target organs in larval and adult D. melanogasler, or even to potentially block receptors ill vivo in physiological studies on a larger species of fly. While this review has concentrated on the structural identification of DHs and ADHs, their expression in NSCs, mode of action on MTs and hindgut, and physiological relevance for the control of excretion, it should not be forgotten that these NSCs are neurons and therefore can be expected to participate more generally in the complex cascade of events that results in an animal entering a new behavioural state. These behavioural states would be ones associated with events that might compromise salt and water balance such as feeding, high metabolic activities (flight), ecdysis, etc. Thus, the NSCs expressing DHs possess central projections, providing the necessary neural link to the nervous/endocrine systems. In addition, DHs are expressed in a variety of cell types including interneurons and possibly sensory neurons, and so it is feasible, and should be anticipated, that these neuropeptide families (and indeed serotonin) might represent functional units that interact to modulate behaviour. These functional units and the neuroactive chemicals that they release might bias, at many levels and sites, neuronal, hormonal and physiological events towards a new functional state of the animal. Behaviours such as feeding cannot occur in isolation, but must be part of a behavioural sequence with distinct phases. The families of DHs/ADHs are ideally suited to co-ordinate the quite disparate physiological events that are associated with a common behaviour. That said, it is also clear from this review that essentially nothing is known about the neurobiology of the DH/ADH-containing NSC (or other cell types); no concept of the uniquely identifiable D H / A D H cell', no information on the integration of these cells with the nervous/endocrine systems: and no information as to the neuroendocrine circuits leading to their activation, their feedback loops, regulation of synthesis and release, or integration with the peripheral (sensory) system. This is a vacuum that must be filled if we are to understand the true physiological relevance of the active factors and their participation in the varied behavioural states of the insect that ultimately leads to the success of insects. Research into insect DHs and

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A D H s will require the full range o f m u l t i d i s c i p l i n a r y a p p r o a c h e s if we are to unravel their complexities in c o m m u n i c a t i o n a n d have a true u n d e r s t a n d i n g from genomics, t h r o u g h p h y s i o l o g y , to behaviour.

Acknowledgements The a u t h o r s t h a n k the researchers who have c o n t r i b u t e d to the w o r k presented in this review a n d have allowed us to r e p o r t u n p u b l i s h e d findings. W e are grateful to the N a t i o n a l Institutes o f Health and the N a t u r a l Sciences a n d Engineering R e s e a r c h Council o f C a n a d a for their s u p p o r t .

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409

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ADDENDUM The genome sequence of the malarial mosquito, Anopheles gambiae, was published in October 2002 (Holt et al., 2002). Mosquito homologues of the neuropeptides and receptors described in this review are readily identifiable (Riehle et al., 2002; Hill et al., 2002) although, interestingly, to obtain the full sequence of the CRF-like DH another intron/exon excision is needed (J. Vanden Broeck, personal communication) as previously found for Drome-DH (see section 4.2.2,2).

References Hill, C. A., Fox, A. N., Pitts, R. J., Kent, L. B., Tan, P. L., Chrystal, M. A., Cravchik A., Collins, F. H., Robertson, H. M. and Zwiebel, L. J. (2002). G protein-coupled receptors in Anopheles gambiae. Science 298, 176 178. Holt, R. A., et al. (2002). The genome sequence of the malaria mosquito Am)pheles gambiae. Science 298, 129 149. Riehle, M. A., Garczynski, S. F., Crim, J. W., Hill, C. A. and Brown, M. R. (2002). Neuropeptides and peptide hormones in Anopheles gambiae. Science 298, 172 175.

Index Ah('Ls'(m 30 Acanthodis curvidens 239 Acanthogry//us Jbrtipes 248 A('anthoplus speiseri 25 I acetylcholine receptors (AchRs) 114

dcherontia styx dopamine in 99 octopamine in 106

Acheta domesticus Achdo-Dp in 297 CRF-like diuretic hormones in 304, 329 diuretic hormone in 302 dimeric peptide in 293 haemolymph in 379 kinins in 305, 331,361,370 MNCs and LNCs in 358, 364 NSCs in 359 sound signalling in 168, 200, 217, 219. 221,223, 225, 251 synergism between diuretic hormones 380 transport in Malpighian tubules 285 acridid ear 176 80 adipokinetic/red pigment concentrating hormone (AKH) family 293 Aeries aeg;vpti 285, 373 CRF-related peptide in 330, 357 diuretic/myotropic kinin neuropeptides in 305, 33l, 333. 335 haemolyph in 380 kinins in 360, 361,364 MNCs and LNCs in 358, 364 mosquito natriuretic peptide (MNP) in 311 NSCs in 359 post-eclosion diuresis in 290 serotonin in 324, 325, 351 A~amtcris iiTsectivora 239 Agrotis ,s~?gellt171359

A I/omenobius socius 214 A//onemohius ./ilsciatus 214

Ambl)'cor37~ha parvipenni,s" 219, 248 Amelrus 229 1 -aminocyclobutane-trans- 1,3dicarboxylate 73 v-aminobutyric acid see GABA Amphiacusta m(O'a 246 Anahaella 26 ,4nahrus simplex 176 Ancistrura ni~woviltata 167, 201 2, 203, 215 Anopheles gambiae 11, 12, 281 Anostostoma australasiae 156 ANP binding 6 Antheraea pernyi 108 Antheraea polyphemus 27, 40 antidiuretic factors (ADFs) 284 antidiuretic hormones see diuretic and antidiuretic hormones Arltrozous p, pullidus 225

Amtrogryllus 156 Amtrogwllus arboreus 162 Apis melfilera diuretic and antidiuretic hormones and 301, 305,358, 359, 360 dopamine in 98 EAAT (apmEAAT) 64, 67, 70 arginine vasopressin-like insect diuretic hormone (AVP-IDH) 294 6, 326, 351 2, 368

Armadillicfium vulgare 317 aromatic amino acid decarboxylase (AADC) 59 Astacus aslacus, serotonin in 92 atrial natriuretic peptide 5 atropine 121 auditory interneurons 194 206 ascending 201 3 in grasshoppers 205 6 in the mole cricket 203 5 omega neuron 197 201 T-cell 194 7

412

auditory receptor organs in the tibia 182 5

Balanus *mbilis 122 Balboa libialis 239 Barbitistes 212 Barbitistes serricauda 225 Berkeley Drosophila Genome Project 116, 307 Blaberus 96 dopamine in 101, 104 octopamine in 109 Blaberus discoMalis 27 BLAST analysis of Drosophila genome 3, 8, 15, 23, 30, 293, 303 Bombyx, protein kinases in 27 Bomhyx mori 321, 322, 383 guanylyl cyclase in 2, 44 Boophilus microplus 331 Bullacris memhracioMes 160, 181, 211 ce-bungatoxin 116 C. morostts 342, 344 C. salinarius 302, 330 C. vicina 375 Caenorhahditis eh',gans 126 choline transporters 116 dopamine in 102 MsGC-fi3 21 MsGC-I 18 octopamine in 107 receptor GCs in 3, 34 calcitonin-like peptides 304, 336, 363 (2dliphora ervthrocephala 92, 349 Calliphora vomitoria 358, 359, 360 calyculin A 39 Cancer borealis 88 Carcinus maenas 70, 92, 317 cardioacceleratory peptide 2b (CAP2b) 42 3,307 8, 336, 346-7, 362 3, 37l Ceratitis capitata, protein kinases in 27 cGMP-dependent protein kinase (PKG) 1 chloride transport stimulating hormone (CTSH) 314--15 Choeroparm~ps 239 choline acetyltransferase (CHAT) 58 choline transporters 114~21 background 114~15 distribution 119 kinetics and pharmacology 119 21 regulation 121 structure 116 19

INDEX

Chomh'oderella borneenses 240 Chortkippus 214 Chorthippus h~uttulus 165, 179, 180, 187, 188, 189, 191, 192, 193,215 Chorthippus mollis 164 choruses 247 51 alternating 248 synchronous 247 unison bout singing 247 unison singing 247 Ckymomyza costata, dopamine in 101 Ciona intesthutlis, MsGC-I 18 cis-3-aminocyclohexanecarboxylic acid 87 (2S,3S,4R)-cis-(carboxycyclopropyl)glycine (CCG III) 73 "clockwork cricket" 161 Owmidophyllum e.vimium 228 cocaine 109, 110 Conocephahls 246 Conocephalus brevipemfis 214 Conocwhalus conocepkalus 235 Comwephalus maculatus 235 Conocephalus n&ropleurum 214, 218, 245 Copiphora 239 Copiphora hrevirostris 238, 239 corticotropin releasing-factor see CRF CRF-related diuretic peptides 293, 327 mode of action 329 1 receptors 326 CRF-related neuropeptides 296 304 isolation and purification 296 302 structures of CRF-related DH 3 0 3 4 CRF-related peptides 344~5, 352 7 circulating levels 368 70 degradation and inactivation 375 6 crustacean cardioactive peptide (CCAP) 21, 37, 293 crustacean hyperglycemic hormone (CHH) 8 cyclic GMP l 44 ecdysis 37 41 f o o d - s e a r c h b e h a v i o u r 41 2 function 26 43 Malpighian tubule regulation 42 3 molecular targets 26-32 cyclic nucleotide-gated channels 30 2 protein kinases and substrates 27 30 neuronal development 34-7 physiological function 32~43 regulation 2 26 sensory physiology 33~4

INDEX

cyclic nucleotide-gated channels 30 2 cyclic nucleotide-gated ion channel protein (cng) 30 cny#like (cn~/) 30 CvcloptiloMes canariensis 160, 161 Cvphoderris 156 Qwhoderris monstrosa 181, 186, 243, 244, 246 CyphoderrLs strepitans 221. 244

D-cysteate 73

Dectk'us verrucivorus 183 Deinacrida 157, 228 Deinacrida rugosa 229 desipramine 104, 105 Dictostelium, atypical guanylyl cyclases 15 Diploptera punctata 293, 294, 301, 380 EAAT (dipEAATI) 64 diuretic and antidiuretic hormones 279 388 cellular actions 324- 47 co-localisation 363-6 distribution 348 66 diuretic/myotropic kinin neuropeptides 331 5 calcitonin-like peptides 336 CAP?b/PVK-2 336 mode of action 333 5 partially characterised factors acting on Malpighian tubules 337 8 receptors 331 3 Tenehrio ADFoe (Tenmo-ADFoe) 337 fluid uptake from the cryptonephric complex 341 2 functions 289 91 clearance of toxic wastes 291 excretion of excess metabolic water 290 post-eclosion diuresis 290 postprandial diuresis 289 90 restricting metabolite loss 291 integrated activities 378 83 co-ordinating Malpighian tubule/ hindgut activities 382 3 excretory system as target for pest control strategies 383 5 future directions 385 7 haemolymph volume/composition maintenance 378 80 synergism between diuretic hormones 38/t 2

413

isolation/structural characterisation of active factors 291 324 in neurosecretory cells and neurohaemal structures 348-66 physiological relevance 366 77 purification and chemical structure of neuropeptides that act on Malpighian tubules 293 5 that stimulate locust hindgut 312 24 regulation of hindgut activity 338 40 of Malpighian tubule secretion 324 31 structure/activity studies 342 7 diuretic/myotropic kinin neuropeptides 305 7, 331 5 DocMocercus 239 DocMocercus gigliotosi 239 DOPA decarboxylase (DDC) 58 dopamine (DA) 91, 110 dopamine transporter (DAT) 58, 99 106 background 99 102 distribution 103 functional domains 102 3 glycosylation sites 103 kinetics and pharmacology 103 5 phosphorylation sites 102 regulation 105 6 st.ucture 102 3 Drepanoxiphus an~ustekmfinaltts 239

Drosophih* aspartate transporter (DrmEAAT2) 68, 69, 77, 78 atypical GCs in 22 Blotgene 112, 113 CG17922 gene 311-1 CG3536 gene 30 1 cn~ 30, 33 DA-ergic neurons in 101 DAT in 103 dopamine in 101, 104 drmDAT 96. 105 dunce (~hlc) 23 ea~ family 31 2 eclosion in 39, 4(/ enahh, d (enh) gene 30 excitatory glutamate in 62 /braging (lbr) gene 41, 42 GABA transporters 80, 86 glial cells in 88 glutamate receptors in 61

414

INDEX

Drosophila (contimwd)

EGPs 29

glutamine cycle 76 histamine in 122 hyperpolarization activated (//h) channels 31, 33 hwbriated (ine) gene 82, 83, 112, 113 hw transporter 112 Malpighian tubule regulation 42 MsGC-I 18, 22 neurotransmitter transporters in 60, 61 noq)A gene 33 octopamine in 106, 109 orphan transporters in 112 phosphodiesterases in 23, 24, 26 photoreceptors 34, 35 protein kinases in 27 PKG in 27, 29, 36 receptor guanylyl cyclases in 3 11 roho (axon guidance receptor) 30 Rosa gene product 112 rosA mutant 113 serotonin in 92 serotonin transporter (drmSERT) 93, 96 soluble guanylyl cyclases in 11-15 Drosophila Genoma Project 2

Elephantodeta nobilis 249

Drosophila mehmogaster calcitonin-like peptides 304 CAP?~ in 308, 336, 346, 361, 362 CRF-related diuretic peptides in 327, 330 diuretic/myotropic kinin neuropeptides in 331,333, 335 dopamine in 102 Drome-DH31 291,387 EAAT (drmEEATs 1 and 2) 64, 67, 70 GABA transporters 79 genome 126, 281,296, 303,386 ITP sequence 321,322, 323 kinins in 357 358, 359, 360, 377 Manse-DH in 383 octopamine in 110 Peram-pyrokinin 5 in 365 serotonin in 92, 325 songs in 222 V-ATPase in 329 Drosophiht punctata 301, 302, 336, 363 D-threo-3-hydroxyaspartate 73 ecdysis 37 41 ecdysis-triggering hormone (ETH) 37 eclosium hormone (EH) 37

Ena/VASP-like protein (EVL) 30 Eneoptera gto,anensis 227 Ephippiger 182, 223 Ephipp~zer ephippiger 155, 165, 167, 169, 222 El)hipl?igeri&~ taeniata 67 Eunemobius carol#ms 235, 245-6

Euphasioptet3,x ochracea (Ormia ochracea) 224, 229, 230, 231,233,241,252 excitatory amino acid transporters 61 129 applications to insect control 125-9 future directions 127 8 neurotransmitter transporters as new targets for 126 7 postgenomic prospects for research 128 9 relevance of insect neurophysiology to 125 Na+-dependent transporters II 121 3 Na-/C1-dependent GABA and monoamine transporters I 78 114 Na +/Cl--dependent transporters II 114 21 N a t / K +-dependent aspartate transporter 77 8 Na + K • -dependent glutamate transporters 61 77 putamine neurotransmitter transporters 123 5 excretion, physiology of 282-8 fluid reabsorption across the cryptonephric complex 288 food-search behaviour 41 2 introduction 282-4 transport processes in hindgut 285 8 in Malpighian tubules 284-5

Formica polyctena 304, 337 Formica polyctena antidiuretic factor (FopADF) 311 12

Formica ru/~l, serotonin in 92 frequenin 10, 18 frog epinephrine transporter (lET) 111

G protein-coupled receptor kinases 10 G. bimaculatus 168 GABA 78

INDEX

GABA transporters (GATs) 78, 79 91 background 79 80 co-localization of EAAT and GAT in glial cells 88 91 distribution 86 EFWER sequence in EL2 83 functional domains 81 6 heptan leucine zipper motif 82 3 ion-permeation site 82 kinetics and pharmacology 86 7 N-linked glycosylation sites 83 6 PKA and PKC phosphorylation sites 83 regulation 87 8 structure 80 6 substrate binding site(s) 82 Gampsoch'is huergeri 194 Gampsocleis graliosa 163, 182, 184, 185, 189 GBR12909 104 GC-activating proteins (GCAPs) 10 11 glial transporter protein 1 (GET-l) 69 gliapse 88 GLUT 62 glutamate decarboxylase (GAD) 59, 79 glycine transporters 124 Grompha~h;rhina portentosa 62 gryllid ear 180 1 Gryllodes supplicans 156 Gry//otall)a /wxadaclyla 235 Grvllotalpa major 156, 167, 248 Gryllomlpa vineae 160 Grvlhts 247 Gryllus himaculatus 101, 162, 164, 189, 197, 210, 212, 219. 220, 252, 348, 359 Grvlhts campestris 159, 161, 180, 197, 200, 201,222, 252 Gryllus,firmus 168, 212, 230 Gryllus./idtoni 167, 230

Grvllus in:eger see Grvllus texensis Grvllus liHeaticeps 168, 169, 221, 223, 241 Grvlhcs pem~sylvanicus 166 Gryllus ruhetts 167, 229, 230, 231,232, 251 Grvllus te.vensis 220, 224, 227, 229, 230, 231, 232, 233, 241,242, 251, 252 guanosine 3'5' cyclic monophosphate see cyclic GMP guanylyl cyclases 2 22 atypical 15 19 biochemical properties 11 15 ligands and activators 8 11 receptor 3 I1

415

sensory receptor 5 sequence analysis 3 8, 11 15 soluble 11 15

lta&'ogryllacrLv 229 Huematobia irrilans 307 Haenschie/hl ecuadorica 176, 239 haglid ear 181 hearing organs, structure of 1711 81 acridid ear 176 80 age, changes with 181 gryllid ear 180 1 haglid ear 181 tettigoniid ear 171 6 Helicoveq)a zea 305, 377, 384 tleliothis virexcens 26. 31 2, 33, 366, 372. 377, 384 Hemiamh'us 228, 229 hemicholinium-3 116. 120 Hemideina 228 Hemideimt crassidens 156 7 Hemisaga denticulata 190 high-affinity glutamate transporters (EAATs) 59 Hirur& 98 histamine (HA) 9 l histamine transporter (HAT) 58, 121 3 background 121 2 distribution 123 kinetics and pharmacology 123 molecular biology 123 histidine decarboxylase (HDC) 58

Homarus americam~s 317 llomorocor37~hus 235 Homotrixa alleHi 166, 230, 232, 233 human nor-epinephrine transporter (hNET) 93 ttyalophora cecr(qfia 23, 27 6-hydroxytryptamine .s'ee serotonin Hvles litleata 294, 301 imipramine 104, 105 inositol trisphosphate receptor (1P3R) associated PKG substrate (1RAG) 29 hlsclra covilleae 235 hlsara elegcols 235 ion transport peptide (ITP) 315 24, 347. 363 amino acid sequence 316 18 expression of 319 2(/ ITP-like (ITP-L) cDNA in locusts 318

416

INDEX

ion transport peptide (ITP) (continued) purification, partial sequencing and actions 315 16 sequence evolution among insects 3 2 0 ~ synthetic 318 Ischnomela pulchripennis 238 juxtamembrane hinge 6

Kawanaphila mir& 175 Kawanaphila nartee 175-6, 194, 216, 219, 246

Kawanaphila yarraga 175 kinase-like domain 6 kinins 345 6 circulating levels 370 l in neurosecretory cells and neurohaemal structures 357 61 Laupala 214, 215 Laupahl cerasina 214 L-cysteate 73

Leptinotarsa decemlineata 308 10, 348 Leptophyes punctatisshna 155, 220, 223, 244, 248

Lerneca jilscipennis 227 8 Leucophaea, excitatory glutamate in 62 Leucophaea maderae 305, 308, 358, 359, 360, 361, 364, 370 L-glutamate 62 Libanasidus vittatus 157 L~gurotettix coquilletti 218, 244, 250 Ligurotettix phmum 217, 246, 250 Limulus 122 Limulus polyphemus 117

Locusta mi,~,raloria 8 arginine vasopressin-like DH in 295 AVP-like immunoreactive neurons in 351, 368 calcitonin-like peptides 304, 336 CAP2b in 308 choline transporters 116, 121 co-localisation 364, 365, 366 CRF-related diuretic hormone 301 CRF-related diuretic peptide 352, 355, 357 diuretic/myotropic kinin neuropeptides in 331 dopamine in 99, 101 GABA transporters 79 haemolymph in 374 histamine in 123

ion transport peptide 363 kinins 358, 359 Locmi-DH in 297, 384 neuroparsins 312, 313, 314 octopamine in 110 serotonin in 92, 93, 324, 348, 349 sound signalling 225 taurine in 124 L-threo-3-hydroxyaspartate 73

L-trans-pyrroliginre-2,4-dicarboxylate (LPDC) 73

Lymnaea sta~nalis 306, 331 Manduca 96 atypical guanylyl cyclases 15, 17, 18, 19 20, 21 CNS 23 ecdysis 37, 40 EGPs in 29 MsGC-I 17, 18 neuronal development 34~5 orphan transporters in 113 protein kinases in 27 receptor GCs 3 soluble guanylyl cyclases in 11 15 VNCs 38 Mamhlca sexta 70, 372, 386 antidiuretic factors 310 11 CAP~b in 361, 362 cardioacceleratory peptides (CAPs) in 307, 371 co-localisation in 364, 365 CRF-like DH receptors 304 CRF-related peptides 352, 355, 376 cyclic AMP production 345 diuretic/myotropic kinin neuropeptides in 305 fluid uptake from the cryptonephric complex 341 GABA transporters 79, 80, 86 glutamate uptake 62 guanylyl cyclase in 2 histamine in 121, 123 kinins in 358, 359 Manse-DH 295, 296, 300--1 Manse-DPll in 366 octopamine in 106, 108, 109 orphan transporters in 113 receptor GCs in 4 serotonin in 92, 348

Mehmoplus sanguinipes 217 Metaballus litus 190

INDEX

Micronycteris" hirsuta 234, 238 Miogryllus 156 mosquito natriuretic peptide ( M N P ) 311 Motttweta isolala 228 MsGC-fi3 19 22 MsGC-1 17-19 MsGC-II 3 MULTICOIL program 12, 22

Musca GABA transporters 80 histamine in 122 Musca domeslica 77, 297, 299, 336, 346, 352. 355, 358, 359, 360, 364, 377 muscarinic ACH receptors 115-16 Mygalopsis rearM 182, 185. 244 Mygal~q~sis paul~erculus 190 Myopophyllum speciosum 166, 238, 239 Myoti~ mvotis 237 Myrme/eotettix macu/atus 222 Na ' -dependent transporters !I 121 3 Na-/C1--dependent GABA and monoamine transporters 1 78 114 Na ~/C1--dependent transporters il 115 21 Na - K ~ -dependent glutamate transporters 61 77 chloride channel domain 69 dihydrokainate (DHK) binding site 69 distribution 69 72 functional domains 67 9 glutamine cycle 76-7 histidine "326" 69 kinetics and pharmacology 72 4 N-linked glycosylation sites 69 Na ~ binding sites 68 in permeation site 67 8 PKA and PKC phosphorylation sites 68 regulation 74 5 structure 63 9 substrate selectivity domains 67 zinc-binding site 68 Na +-dependent aspartate transporter 77 8 natriuretic peptide clearance receptor (NPR-C) 7 Nauphoeta cinerea 358, 359, 364 Neobellieria bullata 307 Neoconocephalus 169, 245 Neoconocephalus caudellianus 247 Neoconoc~7~halus ensiger 195 6, 207, 229, 235. 236

417

Neoconocephalus Neoconocephalus Neoconocephalus Neoconocephalus

exiliscanorus 247 nebrascensis 247, 249 rohusms 169 spiza 217, 250

neuroealcin 10 neurolls

714 205 6 molecular chemistry 56 9 neurotransmitter uptake and vesicular storage 59 61 neuroparsins 312 14 neurotransmitter receptors 56 neurotransmitter transporters (NTTs) 56, 60 nicotinic receptors (nACHRs) 114 nipecotic acid 87 nisoxetine 104, 105, 109 nitric oxide (NO) 11

nitric-oxide (NO)-insensitive soluble guanylyl cylases (GCs) 2 nitric oxide synthases 1I Nyclophilus geq~/i'o3i 240 Nyctophilus mq/or 240

octopamine (OA) 78, 91, I10 octopamine transporters 106 11 distribution 208 kinetics and pharmacology 108 9 regulation 109 10 structure 10Z 8 tyramine transport 110 11 O D Q (1 lt-[1,2,4]oxadiazolol[4,3-a]

quinoxalin- l-one) 14

Oecanthus 214 Oecanthus celerinictus 162 Oecanthusjidtoni 247, 249, 250 Oecanthus n~,,ricornis 219 Oecanthus quadriptmctatus 162 okadaic acid 42 omega neuron 197 201 in acridids 201 1 (ON1) 197-200 2 (ON2) 200 1 Omocesttts viri~hdus 164, 165, 245 Oncopeltu.s' jasciatus 365 Onymacris p[ana 291 Orchelhmcm 167 Orchelinnm7 gladiator 248 Orche/imum n~wipes 248, 251, 252 Orche/imum vulgare 246, 248 Orconectes ]imosus 363

418

Ormia ochracea ( Euphasiopteryx ochracea) 224, 229, 230, 231,233,241. 252 Orocharis luteolira 230 orphan transporters 78-9, 111 14 background 111 distribution 113 kinetics and pharmacology 113 14 structure 112 13 Otus stops 225 1 H-[1,2,4]oxadiazolo[4,5-a]quinoxalin- 1one (ODQ) 20 3-N-oxalyl- L-2,3-diaminopropionate 74 oxotremorine 121 Pachnoda simuata 322 Paramecium, atypical guanylyl cyclases in 15 Parascopioricus exarmatus 228 Penaeus vannamei 306 Periplaneta americana choline transporters 115 co-localisation in 365 CRF-like diuretic hormone in 302 dopamine in 99, 101 2 GABA transporters 79, 80 glutamate uptake 62 Manse-CAP2wlike 308 NSCs in 359 PerampDP in 297 serotonin in 92, 348 taurine in 124 periviscerokinins see cardioacceleratory peptide 2b Phalacrocera replicata 359, 361 Pllaneroptera./~dcata 237 Phaneroptera nana 219, 248 Pholidoplera griseoaptera 185, 186, 225, 227, 248, 250 Phormia terraenovae 358. 359, 360 phosphodiesterases (PDEs) 22-6 Phv//omimus inversus 240 Pieriv hrassicae 290 Pieris' rapae 296. 345. 372 piperidenecarboxylic acid 88 Plagiostira all)onotata 235 Phtsmodium, atypical guanylyl cyclases in 15 Plal3"cleis. a/l)opunctata 225 Phttvcleis intermedia 247, 251 Platvslolus ohvius 155 Poecilimon 174, 254 Poecilimon a[finis" 215, 232

INDEX

Poecilhnon artedentatus 232 Poecilhnon mariannae 224. 232 Poecilimon nobilis 232 Poecilimon ornatus 216, 223 Poeci/hmm proprinquus 232 Poecilimon schmidti 165 Poecilh~wn thessalicus 241 Poecilimon veluchianus 232 Polysarchus denticauda 171, 172, 173, 174, 182, 183 proctolin 293 Promeca perakana 240 Promeca sumatrana 240 PROSITE analysis 26 Protein Family (Pfam) databases 26 protein kinases and substrates 27 30 Psammodronlus a]?irus 224 Psorodonotus il[yricus 182 Pterophylla beltrani 228 Pterophylla camelE/blia 228, 248, 250 putamine neurotransmitter transporters 123 5 Ramsay assay 282 rat dopamine (rDAT) 93 rat serotonin transporters (rSERT) 93 recoverin I0 Requena verticalis 168, 169, 174, 216, 219, 221 Rhodniusprolirus 125. 371 2, 373, 386 AVP-like immunoreactive neurons in 351 calcitonin-like peptides 304, 363 cardioacceleratory peptide 2b (CAP2b) in 43, 336 co-localisation 364, 365 CRF-related neuropeptides 302. 352, 355, 357, 376 diuresis in 282 dimetic/myotropic kinin neuropeptides in 305. 335 haemolymph in 374, 375, 379. 380 kinins in 358 Manse-CAP2b in 337 Malpighian tubule transport 285 metabolite loss, restricting 291 NSCs in 359, 360 serolonin in 91. 324, 325, 342, 344. 350, 366, 367-8 synergism between diuretic hormones 380, 381 rhodopsin kinase 10

INDEX

419

Ruspolia nitffUk~ 163 4 Ruspolia 235 Ru,v~olia df[lbrens 171, 182, 183, 184 S100B 10

Saccharomyces cerevMae 345, 386 Scapteriscus abhreviatus 203,205 Sccq~teriscus acletus 158 Sc~q)teriscus borellii 203 5, 230, 237 SCal)leriscu.s' didact),lis 235 Scapteriscus vicim~s 158 Schedocentrus 239 Schistocerca americana 301,358,359, 361. 362

Sch&tocerca gregaria (SgITP) 8 antidiuretic factors in 310 arborisation in 351 directional hearing in 187, 188 dopamine in 99 excretion in 284 GABA transporters 79.80 guanylyl cyclases in 22 hindgut activity 338 histamine in 122 1TP sequencing in 323, 363 neuroparsins in 313, 314 postprandial diuresis in 290 serotonin in 92 Sciara,s'aga quadrata 166, 190, 211,230. 231. 232, 233,234 Scopiorinus jra~ilus 228

Scudderia curvicauda 219 "selfish herd' effect 158 serotonin (5-hydroxytryptamine: 5-HT) 59, 78, 91, 110 circulating levels 366 8 degradation and inactivation 374 5 mode of action 325 6 in neurosecretory cells and neurohaemal structures 348 51 receptors 324-5 secretion by Malpighian tubule 324 6, 342 4 serotonin transporter (SERT) 59, 91 9 background 92 3 cocaine binding site 96 distribution 97 functional domains 94 6 heptan leucine zipper 94 ion permeation site 94 kinetics and pharmacology 98 9

monoamine-binding site(s) 94 regulation 99 structure 93 6 tricyclic antidepressant interaction site 94 6 SKF-89976A 87, 88 sound signalling in Orthoptera 151 254 analysis 189 207 auditory intemeurons 194 206 ascending 201 3 in grasshoppers 205 6 in the mole cricket 203 5 omega neuron 197 201 T-cell 194 7 auditory receptor organs in the tibia 182 5 components 209 cooperation/competition between males 243 52 choruses 247 51 satellite males and silent searching 251 2 spacing, aggregating and fighting 244 6 defences against acoustically orienting predators 226 9 against bats 234 40 against parasitoids 229 34 directional hearing 187 9 environment, effects on 209 I1 hearing and ears 169 89 heterospecific sounds 224 43 information content 207 24 mate choice 217 24 mate location 215 17 mating systems 154 9 patterns in calling 157 9 variation in 154 7 new directions 252 3 predator avoidance mechanisms, evolution of 242 3 primary afferents in acridids 186 7 in the prothoracic ganglion 186 sex recognition 215 sexual vs natural selection 241 2 songs and signals 159 69 analysis 190 3 changes with age 165 6 energetic costs of calling 168 9 intensity, distance and size 160 I

420

sound signalling in Orthoptera (continued) songs and signals (continued) mechanisms of sound production 161- 3 pattern generation in crickets 163 5 sex differences 167 temperature effects 167 8 vibratory communication 166 7 species recognition 212 15 structure of hearing organs t70-81 acridid ear 176-80 age, changes 181 gryllid ear 180-1 haglid ear 18 I tettigoniid ear 171 6 symmetry and asymmetry 206 7 tonotopic organization of receptor projections 186 7 of sense cells 183 5 Spodoptera 32 Spodoptera littoralis 106, 108 Spodoptera littorina 358 Steirodo, careovirgulatum 228 Stenobothrus lineatus 247 Steropleurus nohrei 155 Steuropleurux stali 155, 224 Stomoxys calcin'ans 297, 299, 307 swiss cheese (sws) gene 26 7 synaptotagmin 35 Syrbula admirubilis 247 Syrbula./itscovittata 247 taurine transporters 124 5 Teleogt3'lhc~ commodus 207, 213 Te/eogryllus oceanicus 185, 199, 200 -1, 203, 213, 218, 223, 231, 233, 236, 239, 240. 251, 252, 335 Tenebrio ADFc~ (Tenmo-ADF~) 337 Tenebrio molitor 63, 288, 294, 299-300. 304, 308 10, 337, 341 Tetrahymena 15 Tettigonia cantans 166, 183, 213, 235, 245 Tett~onia viridissima 172, 173, 186, 196 8,200, 201,203, 206, 207, 211, 213, 225, 235, 236, 244

INDEX

tettigoniid ear 171 6 Therohia h,onidei 225, 232, 241 tiagibine 88 transmembrane o~-helical domains (TMDs) 63 Trichoplusia ni DAT in 103 dopamine in 102 EAAT (trnEEAT1) 64, 71 GABA transporter 86 glutamate in 91 octopamine in 107 orphan transporters in I13 serotonin transporter in 96 taurine transporter in 125 TRP (transient receptor potential) 32 TRl°-like channels 32 tryptophan hydroxylase (TPH) 59 Tympanophyllum arcu/blium 240 tyramine (TA) 78, 91 tyramine/~-hydroxylase (Trill) 58 tyrosine hydroxylase (TH) 58 vasodilator-stimulated phosphoprotein (VASP) 3O vesicular acetylcholine transporters (yAChT) 60 vesicular excitatory amino acid transporters (vEAATs) 60 vesicular inhibitory amino acid transporters (vIAATs) 60 vesicular monoamine transporters (vMATs) 60

X81IOI)IlS oocytes 72, 98 orphan transporters in 112, l 14 spinal neurons 36 Xenopus laevis 327 Xestoptera cornea 228 xylamine 105 zaprinast 42 Zoolerntopsis nevadensis 294, 300, 301

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