Distribution of acetylcholine receptors in the ... - Wiley Online Library

THE JOURNAL OF COMPARATIVE NEUROLOGY 334~47-58 (1993)

Distribution of Acetylcholine Receptors in the Central Nervous System of Adult Locusts BEULAH LEITCH, BRANWEN L. WATKINS, AND MALCOLM BURROWS Department of Zoology, University of Cambridge, Cambridge CB2 3EJ, England

ABSTRACT A polyclonal antibody raised against nicotinic acetylcholine receptor protein from purified locust neuronal membrane was used to analyse the distribution of antigenic sites within the central nervous system of adult Schistocerca gregaria. Light microscopic examination showed that all principal neuropiles in the thoracic ganglia label with the antibody but that the major tracts and commissures do not. Analysis of this pattern of staining in the electron microscope reveals that the receptor is present on specific synaptic and extrajunctional neuronal membranes in the neuropile. Antigenic sites are also evident on the plasma membranes and within the cytoplasm adjacent to Golgi complexes of some neuronal somata, suggesting that these neurones synthesise nicotinic acetylcholine receptors. In addition to neuronal labelling, there is evidence that the receptor is also present on the membranes of three types of glial cells. The implications of this pattern of receptor distribution are discussed. o 1993 Wiley-Liss, Inc. Key words: neurotransmitter-receptorCNS, immunocytochemistry, locust

Evidence for the importance of nicotinic acetylcholine receptors (nAChRs) in neuronal function in the central nervous system (CNS) of vertebrates (Kelly and Rogawski, '85) and invertebrates (Leake and Walker, '80; David and Sattelle, '84) is increasing. Relatively little is known, however, about neuronal nAChRs compared to muscle nAChRs, partly because their study has been hindered by a lack of suitable probes. Recently, specific antibodies to putative neuronal nAChRs have been developed. Most of these antibodies, however, have been raised against vertebrate peripheral nAChR from mammalian muscle (Tzartos et al., '83) and the electric organs of fish (Tzartos and Lindstrom, '80; Tzartos et al., '811, because these tissues, unlike vertebrate CNS, are rich sources of nicotinic receptor. Insect CNS, in contrast, has a high concentration of nAChR and so offers a source of receptor from central neuronal synapses. A polyclonal, monospecific antiserum has been raised against purified neuronal nAChR protein from insects (Breer et al., '85).The receptor was probed by u-bungarotoxin (a-BTX) binding and isolated from detergent-solubilised ganglionic membranes of locusts. The purified receptor protein migrates as a single band on polyacrylamide gel and cross-reacts with some monoclonal antibodies against Torpedo receptor (Breer et al., '85). It also forms functional receptorichannel molecules when reconstituted into artificial bilayers (Hanke and Breer, '86, '87). The antibody has been used to demonstrate specific labelling of areas of synaptic contact in the neuropile of a O

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number of insect species including the mesothoracic ganglion of the locust (Breer et al.,'85), the terminal abdominal ganglion of the cockroach (Sattelle et al., '891, and the brain of the honeybee (Kreissl and Bicker, '89). In the honeybee, antibody binding in the neuropile coincides with acetylcholinesterase (AChE) sites of activity, which the authors interpret as indicating that the antibody cross-reacts with the bee ACh receptor. Antigenic sites have also been demonstrated in the cytoplasm of neuronal cell bodies in the honeybee (Kreissl and Bicker, '89) and the cockroach (Sattelle et al., '89). The demonstration of immunocytochemical labelling in these insects has, however, been limited to light microscopical sections. In the present study, we analyse the localisation of anti-nAChR antibody binding sites within the thoracic ganglia of adult locusts at both light- (LM) and electron microscope (EM) levels. We show specific labelling of synaptic and extrajunctional neuronal membrane and demonstrate the presence of glial AChRs.

MATERIALS AND METHODS LM immunocytochemistry Adult Schistocerca gregaria (Forskgl) were taken from our crowded culture. The thoracic nervous system was fixed Accepted March 22, 1993 Address reprint requests to B. Leitch, Department of Zoology, University of Cambridge, Downing Street, Cambridge CB2 3EJ, England.

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in situ for 10-15 minutes with 4% paraformaldehyde in 0.05 M phosphate buffer (pH 7.5) containing 0.2 M sucrose. The pro-, meso-, and metathoracic ganglia (n = 24) were dissected out and fixed for a further 2 hours in the same solution before being washed in phosphate buffer overnight. The ganglia were then dehydrated and embedded in wax. Sections (10 ym thick) were cut and stained by use of a modification (Bishop and O'Shea, '82) of the peroxidase/ antiperoxidase (PAP) method of Sternberger ('74). Locust nAChR antiserum was generously supplied by H. Breer. Details of its preparation and specificity are given in Breer et al. ('85). The antiserum was used at dilutions of 1:500 and 1:1,000. Goat anti-rabbit IgG and rabbit PAP were obtained from ICN Immuno-Biologicals and 3,3 diaminobenzidine (Sigma) was used as a substrate for the peroxidase reaction. Sections were photographed with a Zeiss Axiophot microscope.

EM immunocytochemistry Ganglia were initially fixed in situ in 5% glutaraldehyde in 0.1 M phosphate buffer (pH 7.2) containing 0.2 M sucrose for 10 minutes. The ganglia were then removed, bisected and fixed for a further 1-2 hours at room temperature in the same fixative. After several washes with phosphate buffer, the tissue was treated with 1%osmium tetroxide for 30 minutes and stained en bloc with 2% aqueous uranyl acetate. The tissue was subsequently dehydrated in graded ethanols and embedded in LR White resin. Sections were cut on an LKB V ultramicrotome with a diamond knife and mounted on nickel slot grids coated with Formvar. LR White sections were etched for 5-20 minutes with saturated sobum metaperiodate (Bendayan and Zollinger, '83).After thorough washing with distilled water, the grids were floated, section face downwards, on droplets of 5% normal goat serum in bovine serum albumin (BSA)/Tris buffer (pH 7.4) for 30 minutes. The grids were then transferred to droplets of rabbit anti-AChR antiserum in BSA/Tris buffer (pH 7.4) for 2-3 hours at room temperature. The antibody was tested at doubling dilutions from 1 5 0 to 1:2,000. Optimal staining was achieved with dilutions in the range 1:lOO-1:200. After a series of at least 5 washes in BSAiTris buffer, the grids were transferred to droplets of 10 nm gold-labelled goat anti-rabbit antibody

(Janssen) diluted 1:20 in BSA/Tris (pH 8.2) for 1 hour at room temperature. The grids were then washed in a series of distilled water droplets to remove any unbound gold conjugate and stained with uranyl acetate and lead citrate (3 minutes each). Control sections were treated identically except for omission of the primary antibody and its replacement with buffer.

RESULTS LM localisation of anti-AChR antibody binding sites All the thoracic ganglia show the same pattern of staining when labelled with the anti-AChR antibody. Neuropiles are stained but tracts and commissures are not (Fig. 1A-E). The principal neuroanatomical features of the thoracic ganglia have been described by Tyrer and Gregory ('82) and Pfliiger et al. ('88) and we use their terminology.

Principal neuropiles The principal neuropiles, where synaptic interactions occur between the various types of neurone, comprise areas of densely packed, small-diameter processes that stain with the anti-AChR antibody. Control sections, treated identically but with omission of the primary antibody step, show no staining. No differences could be detected at the light microscopical level in the density of staining in the different areas of neuropile (Fig. 1). For example, a prominent ventral region of neuropile called the ventral association centre WAC) receives projections from mechanosensory neurones associated with the hind legs, that may use acetylcholine as their transmitter (Callec, '74; Lutz and Tyrer, '88; Sattelle and Breer, '90). The processes here are thin, closely packed, and stain uniformly with the antibody. The dorsal neuropiles, by contrast, contain predominantly the processes of many motor neurones and interneurones, and only the most lateral regions, called the anterior (aLAC) and posterior (pLAC) lateral association centres which contain terminals of some proprioceptive afferents (Pfluger et al., '88) are as densely packed as the ventral neuropiles. Nevertheless, they stain uniformly with the antibody.

Tracts and commissures Abbreviations

AChR CT DC I-VI DIT DMT FETi IT LDT LAC LVT MDT MVT PVC RT SMC

TT VAC VC I VCLII VIT VLT VMT

acetylcholinereceptor C-tract dorsal commissures I-VI dorsal intermediate tract dorsal median tract fast extensor tibiae motor neurone I-tract lateral dorsal tract lateral association centre lateral ventral tract median dorsal tract median ventral tract posterior ventral commissure ring tract supra median commissure T-tract ventral association centre ventral commissure I ventral commissure loop I1 ventral intermediate tract ventral lateral tract ventral median tract

In contrast to the neuropiles, at the LM level of resolution, none of the longitudinal tracts containing bundles of axons appear to stain with the antibody (Fig. 1). For example, the median dorsal tract (MDT); the lateral dorsal tract (LDT); the dorsal intermediate tract (DIT); the ventral intermediate tract (VIT); the dorsal median tract (DMT); the ventral median tract (WIT); the ventral lateral tract (VLT); the median ventral tract (MVT), and the lateral ventral tract (LVT), appear unstained. Similarly, the four major vertical and oblique tracts: the T-tract (TT); ring

Fig. 1. A-E: Transverse light microscopic (LM) sections through a metathoracic ganglion, labelled with the anti-AChR antibody, at the levels indicated on the drawing. All the principal neuropiles are evenly stained with the antibody. In contrast, none of the major longitudinal and vertical tracts or commissures are labelled. Only the major anatomical landmarks are indicated. For detailed descriptions of the major neuropiles and tracts see Tyrer and Gregory ('82) and Pfluger et al. ('88). See Abbreviations list for complete descriptions. Scale bars = 100 pm.

Figure 1

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50 tract (RT); C-tract (CT); and I-tract (IT), also appear t o be unlabelled. None of the major commissures, including the 6 dorsal (DCI-DCVI), and 4 ventral ones [ventral commissure I (VCI); ventral commissural loop I1 (VCLID; supramedian commissure (SMC); and posterior ventral commissure (PVC)] stain with the antibody at this level of resolution (Fig. 1).

EM localisation of anti-AChR antibody binding sites Neuropile. At the EM level, the binding sites of the anti-AChR antibody within the neuropile are located, according to the distribution of gold particles, predominantly on the plasma membranes of neuronal processes (Figs. 2, 3). Membrane staining is extensive throughout the neuropile, but little gold is present over the cytoplasm or organelles of neuronal processes (Fig. 3). The antibody binds to both synaptic and extrajunctional membrane (Figs. 3, 4).At immunopositive synapses (Figs. 3 4 , gold particles are peppered along the synaptic membrane density. The gold label, however, is generally no more concentrated a t positive synaptic sites than it is at positive extrajunctional sites. There is, nevertheless, selective labelling of some synapses (compare Figs. 5 and 6 ) and of the extrajunctional membrane of some processes. For example, in Figure 5, a labelled dyadic output synapse is present in which both postsynaptic profiles have gold associated with their extrajunctional membranes. In contrast, the postsynaptic profiles at the unlabelled synapse in Figure 6 have no gold associated with their extrajunctional membranes. Processes in which several output synapses are evident in a single section, and which all are labelled with gold particles, are assumed to be cholinergic (Figs. 3, 4). Conversely, processes with exclusively unlabelled output synapses (Figs. 2B, 6) are presumed to be noncholinergic. For example, the process in Figure 2B makes three output synapses, all of which are negative for AChR, and receives one input synapse, which is positive. The interpretation is that this process has receptors for ACh but does not release ACh at its own synaptic terminals. Major tracts. The cytoplasm in the axons of the tracts shows no labelling at the EM level (Fig. 7A-C), thus explaining why the tracts appear unstained in LM sections (Fig. 1B-F). There is, however, some labelling of axonal membrane, which is only detectable by ultrastructural examination (Fig. 7B). Neuroglia. The membranes of a t least 3 types of glia (classified according to Strausfeld, '76) also stain with the antibody. First, gold particles are evident along the seams of glial processes that envelope the axons. They overlie the membranes of this type of neuroglia but not the glioplasm or the microtubules, which are the main structural elements (Fig. 7C). Second, glia between the neuropile and the cortical layer of cell bodies (Fig. 8A) have gold particles associated with electron-dense bodies within their cytoplasm (Fig. 8B). Third, the membranes of neuroglia that extend from the perineurium around the apical region of the neuronal somata to form the trophospongia also stain (Fig, 10A,B). The neural lamella and perineurium, however, are unstained. Somata. Neuronal somata have antigenic sites within their cytoplasm. At the LM level, these appear as positively stained granules (Fig. 9). In EM sections (Figs. 10, II),

these granules label with gold particles and are adjacent to Golgi complexes and to smaller immunopositive vesicles (Figs. 10B, 11B). The plasma membrane of the neuronal somata is also labelled discontinuously with gold particles. The nuclear cytoplasm and nuclear membrane, however, show no labelling (Fig. 10B).

DISCUSSION a-BTX as a neuronal nAChR probe The antibody used in this study was raised against purified nAChR as probed by a-BTX binding. a-BTX binding has also been used in other studies to isolate AChR from the supraoesophageal ganglion of locust Schistocerca gregaria (Filbin et al., '83). It is uncertain whether a-BTX binding sites represent functional receptors. Could the antibody be detecting antigenic regions in the insect CNS, which are non-AChR, toxin binding sites? Many studies show that although a-BTX binds to central and peripheral sites with high affinity (Morley and Kemp, '81; Oswald and Freeman, '81)?it often does not block functional transmission at central or peripheral cholinergic synapses in either vertebrates or invertebrates (for review see Schuetze and Role, '87). Furthermore, in mammalian CNS, there is little correspondence between the regional distribution of radiolabelled nicotine and acetylcholine and that of a-BTX (Marks and Collins, '82; Clarke et al., '84, '85). There are, however, exceptions to the general observation that a-BTX does not block neuronal AChR function. The toxin has been shown to block AChR function in frog ganglia (Marshall, '811, and in the superchiasmaticnucleus of the hypothalamus and in the inferior colliculus (Zatz and Brownstein, '81; Farley et al., '83).It also blocks function in the following invertebrates: leech (Pellegrino and Simonneau, '84); Aplysia (Kehoe et al., '761, and, of especial relevance for the present study, insects such as locust (Sattelle et al., '80, '83; Breer et al., '85; Benson, '92). Furthermore, several autoradiography studies have demonstrated that 1251-a-BTXis localized at binding sites in the neuropile of several insect species (Schmidt-Nielsen et al., '77; Hildebrand et al., '79; Meyer and Reddy, '85). Hence in these animals, a-BTX may indeed be a useful marker for neuronal AChRs, and thus its use to probe for ACh receptor protein for the generation of this polyclonal antibody is justified. Ultimate proof that the purified membrane protein is not only a toxin binding site but also a functional receptor is demonstrated when purified receptor is reconstituted into artificial lipid membranes and forms functional receptorichannel molecules (Breer and Sattelle, '87).

Synaptic and extrajunctional antibody binding sites In locust CNS, the anti-AChR antibody binds to both synaptic and extrajunctional neuronal membrane, suggesting that ACh receptors are not localised only at synapses. Receptors for L-glutamate in locust muscle fibres are also present at synaptic and extrajunctional sites (Cull-Candy, '76; Usherwood, 'go), and in cockroach, there is physiological evidence for the presence of presynaptic or extrasynaptic ACh receptors in the membrane of cholinergic axons (Blagburn and Sattelle, '87). In Drosophila CNS, monoclonal antibodies with defined specificities for Torpedo nicotinic acetylcholine receptor label axonal tracts in addition to neuropile regions (Chase et al., '87). By contrast in vertebrates, both central and peripheral AChRs are predom-

DISTRIBUTION OF ACETYLCHOLINE RECEPTORS

Fig. 2. A Electron micrograph of thin sections through a metathoracic ganglion shows antibody binding sites within the neuropile. Gold particles are localised predominantly over the plasma membranes of neuronal processes, with few over the cytoplasm or organelles. B. Highpower view of the region indicated by the box in A. A noncholinergic

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process (asterisk) makes three output synapses (arrowheads), all of which are immunonegative for AChR, and receives one immunopositive input synapse (double arrowhead). Scale bars = 1 +m for A 0.5 +m for B.

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Fig. 3. A probable cholinergic process (asterisk) with three output synapses (S, large arrowheads) that are all labelled with gold particles. Gold is also localised along the extrajunctional (EJ, small arrowheads) membranes of some processes. Scale bar = 0.5 pm.

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Fig. 4. A probable cholinergic process (asterisk) with immunopositive output synapses (arrowheads). Scale bar = 0.5 pm.


Fig. 5. High-power view of two immunopositive dyadic output synapses (S, arrowheads) on a probable cholinergic process (asterisk). The gold is localised over the synaptic membrane density and the extrajunctional membranes of both postsynaptic processes. Scale bar = 0.2 IJ-m.

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Fig. 6. High-power view of two immunonegative dyadic output synapses (S,arrowheads) on a probable noiicholinergic process (asterisk). There is no labelling of the synaptic membrane or of the extrajunctional membrane of the postsynaptic processes. Scale bar = 0.2 Fm.

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Fig, 7. A: Electron microscopic (EM) section through aventral commissure composed of bundles of axons, the axoplasm of which shows no labelling. B and C are high-power views of the regions indicated by the boxes in A. B: Note gold particles along the apposed membranes of two axons. C : Gold is also evident over the membranes of interposed glial processes. Note there is no gold over the glioplasm or the glial microtubules (MT). Scale bars = 0.2 pm for A; 0.5 Fm for Band C.

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Fig. 8. A: EM section shows the glia (G) at the interface between the neuropile (N) and the cortical layer of cell bodies (0. B: High-power view of the region indicated by the box in A. The glial process contains electron-dense bodies, which have gold associated with them. Scale bars = 5 pm for A 1 pm for B.


Fig. 9. LM section of neuronal cell bodies in the ventral region of the ganglion. Antigenic granules (arrowheads) are evident in the cytoplasm. Scale bar = 50 pm. pigs, 1 0 and ~ 1 1 ~ .EM sections through neurond cell bodies show the presence of electron-dense granules in the cytoplasm ofthe somata. Scale bars = 5 pm and 4 pm, respectively.

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Figs. 10B and 11B. High-power views of the regions indicated by the boxes in Figures 10A and 11A, respectively. Gold particles are localised over granules (Gr) and vesicles adjacent to Golgi complexes (Gc), on the plasma membrane (Pm) but not on the nucleus (N). Some gold is also associated with the plasma membrane of the neuroglia (arrowheads) that invaginate into the neuronal somata to form the trophospongia. Scale bars = 1km and 0.4 km, respectively.

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inantly localised at synapses (Peper and McMahon, '72; Matthews-Bellinger and Salpeter, '78; Marshall, '81; Jacob et al., '84). Extrajunctional receptors may have different functional properties from synaptic receptors. For example, endplate channels associated with AChRs in adult frogs differ from those that appear extrajunctionally after denervation (Katz and Miledi, '72). Endplate channels carry larger currents and close more quickly than extrajunctional channels, thus leading to their designation as fast and slow channels, respectively (Schuetze and Role, '87). Both fast and slow AChR channels have been identified in rat submandibular ganglion neurones. It is suggested that the fast and slow channels are spatially separate and that spontaneous transmitter release occurs only at regions where there are fast channels (Rang, '81). In the present study, it is not known whether the extrajunctional and synaptic ACh receptors, identified in locust with this antibody, are associated with channels of different properties. Tareilus et al. ('90) identified two physiologically different agonist-activated channel types in locust CNS, which they supposed represented synaptic and extrasynaptic acetylcholine receptors. The supposed synaptic type showed a high conductance and a short mean lifetime whilst the extrasynaptic type had a lower conductance but a longer lifetime. Two pharmacologically distinct types of extrajunctional L-glutamate receptors also occur in locust muscle fibres (Cull-Candy, '76). In the CNS of cockroach, glutamate applied to extrajunctional receptors on the soma of the fast coxal depressor motor neurone induces a hyperpolarisation (Wafford et al., '911, whereas it causes a depolarisation of motor neurones when injected into synaptic regions of locust neuropile (Sombati and Hoyle, '84; Dubas, '90).

Neuronal somata antibody binding sites AChR-immunoreactive granules are evident in the cytoplasm of some neuronal cell bodies and are associated with Golgi complexes, suggesting that these neurones are active in the synthesis of nACh receptors. For muscle AChR, only complete receptor molecules are transported to the cell surface and then inserted (for review see Schuetze and Role, '87). Each of the AChR subunits is translated from a separate mRNA, inserted into the endoplasmic reticulum and glycosylated. Processing by the Golgi apparatus is inferred by the presence of complex oligosaccharides on the mature receptor. Additional modifications, such as phosphorylation, disulphide bond formation, and methylation may also occur. Finally, the AChR complex is assembled, but the site of this assembly is not known. Antibody binding sites are also evident along the plasma membrane of some of the locust somata. AChRs have been demonstrated pharmacologically on the cell bodies of insect motor neurones (David and Pitman, '82; David and Sattelle, '84) and interneurones (Harrow and Sattelle, '83). In locust, AChRs have been detected on the somata of embryonic dorsal unpaired median (DUM) neurones as early as day 8, which is at approximately 40% of embryonic development, long before any spontaneous synaptic input is seen (Goodman and Spitzer, '80). There is no conclusive evidence, however, that somatic and synaptic receptors are identical.

Glial antibody binding sites The localisation of gold particles along the membranes of particular types of glial cells suggests that receptors for

acetylcholine are present on both glia and neurones. This finding accords well with the presence of various receptors on glial cells in other animals. For example, nicotinic cholinergic receptors have been located autoradiographically on the membranes of Schwann cells in squid (Rawlins and Villegas, '781, and functional receptors for neurotransmitters have been found on astroglial cells in mammals (Murphy and Pearce, '87). Functional nAChRs have also been characterised in leech neuropilar glial cells (Ballanyi and Schlue, '89) and in crayfish glial satellite cells (Lieberman et al., '81). Primary cultures of glia and cell lines of glial origin from central and peripheral nervous system have receptors for catecholamines, some peptides and prostaglandins (for review see Van Calker and Hamprecht, '80). It appears, therefore, that glia possess an array of receptors coupled to ion channels and intracellular signalling pathways that may be very similar and even related at the genomic level, to those present in neurones (Pearce, '91). Furthermore, glial cells are able to accumulate, synthesise, and release neurotransmitters or other neuroactive substances (Villegasand Jenden, '79; Villegas, '84). Activation of these receptors can alter the membrane properties of a glial cell. For example, in crayfish, acetylcholine inactivates C1- channels in satellite glial cells, and thus induces a membrane hyperpolarisation (Brunder and Lieberman, '88). This leads to a K+ uptake, which could buffer K+ during axonal activity. ACh also induces membrane hyperpolarisation in squid Schwann cells (Villegas, '84) and rat glioma cells (Hamprecht et al., '76). In leech, however, ACh and nicotinic agonists depolarise CNS glial cell membrane (Ballanyi and Schlue, '89). The presence of AChRs in locust glial cells indicates that these sorts of effects must now be considered as possible mechanisms in the integrative actions of the neuronal circuits that process sensory signals and generate motor patterns.

ACKNOWLEDGMENTS This work was supported by a grant from the SERC, UK. Many thanks are due to H. Breer for kindly supplying the AChR antibody. We thank W. Lee for printing the photographs and our Cambridge colleagues, and in particular D. Sattelle, for their critical comments on this manuscript.

LITERATURE CITED Ballanyi, K., and W.R. Schlue (1989) Electrophysiological characterization of a nicotinic acetylcholine receptor on leech neuropile glial cells. Glia 2t330-345. Bendayan, M., and M. Zollinger (1983) Ultrastructural localization of antigenic sites on osmium-fixed tissues applying the protein-A-gold technique. J. Histochem. Cytochem. 31t101-109. Benson, J.A. (1992) Electrophysiological pharmacology of the nicotinic and muscarinic cholinergic responses of isolated neuronal somata from locust thoracic ganglia. J. Exp. Biol. 170:203-233. Bishop, C.A., and M. O'Shea (1982) Neuropeptide proctolin immunocytochemical mapping of neurones in the central nervous system of the cockroach. J. Comp. Neurol. 207223-238. Blagburn, J.M., and D.B. Sattelle (1987) Nicotinic acetylcholine receptors on a cholinergic nerve terminal in the cockroach, Periplaneta um.ericana. J. Comp. Physiol. 161t215-225. Breer, H., and D.B. Sattelle (1987) Molecular properties and functions of insect acetylcholine receptors. J. Insect Physiol. 33:771-790. Breer, H., R. Kleene, and G. Hinz (1985) Molecular forms and subunit structure of the acetylcholine receptor in the central nervous system of insects. J. Neurosci. 5:3386-3392.

DISTRIBUTION OF ACETYLCHOLINE RECEPTORS Brunder, D.G., and E.M. Lieberman (1988) Studies of axonal-glial cell interactions and periaxonal K+ homeostasis-I. The influence of Na+, K+, C1- and cholinergic agents on the membrane potential of the adaxonal glia of the crayfish medial giant axon. Neuroscience 25:951969. Callec, J.J. (1974) Synaptic transmission in the central nervous system of insects. I n J.E. Treherne (ed): Insect Neurobiology. Elsevier, New York. pp. 119-185. Chase, B.A., J. Holliday, J.H. Reese, L.L.Y. Chun, and E. Hawrot (1987) Monoclonal antibodies with defined specificities for Torpedo nicotinic acetylcholine receptor cross-react with Drosophila neural tissue. Neuroscience 21:959-976. Clarke, P.B.S., C.B. Pert, and A. Pert (1984) Autoradiographic distribution of nicotine receptors in rat brain. Brain Res. 323t390-395. Clarke, P.B.S., R.D. Schwartz, S.M. Paul, C.B. Pert, and A. Pert (1985) Nicotinic binding in rat brain: Autoradiographic comparison of 125-abungarotoxin. J. Neurosci. 5r1307-1315. Cull-Candy, S.G. (1976) Two types of extrajunctional L-glutamate receptors in locust muscle fibres. J. Physiol. 255r449-464. David, J.A., and R.M. Pitman (1982) The effects of axotomy upon the extrasynaptic acetylcholine sensitivity of an identified motoneurone in the cockroach Periplaneta americana. J. Exp. Biol. 98;329-341. David, J.A., and D.B. Sattelle (1984) Actions of cholinergic pharmacological agents on the cell body membrane of the fast coxal depressor motoneurone of the cockroach (Periplaneta americana). J. Exp. Biol. 108:119136. Dubas, F. (1990) Inhibitory effect of L-glutamate on the neuropile arborizations of flight motoneurons in locust. J. Exp. Biol. 148:501-508. Farley, G.R., B.J. Morley, E. Javel, and M.P. Gorga (1983) Single-unit responses to cholinergic agents in the rat inferior colliculus. Hear. Res. 11:73-91. Filbin, M.T., G.G. Lunt, and J.F. Donnellan (1983) Partial purification and characterisation of an acetylcholine receptor with nicotinic properties from the supraoesophageal ganglion of the locust (Schistocerca gregaria). Eur. J. Biochem. 132151-156. Goodman, C.S., and N.C. Spitzer (1980) Embryonic development of neurotransmitter receptors in grasshoppers. In D.B. Sattelle, L.M. Hall, and J.G. Hildebrand (eds): Receptors for Neurotransmitter, Hormones, and Pheromones in Insects. Amsterdam: ElsevieriNorth Holland, pp. 195207. Hamprecht, B., W. Kemper, and T. Amano (1976) Electrical response of glioma cells to acetylcholine. Brain Res. 101r129-135. Hanke, W., and H. Breer (1986) Channel properties of a neuronal acetylcholine receptor purified from central nervous system of insect reconstituted in planar lipid bilayers. Nature, Lond. 321: 171-174. Hanke, W., and H. Breer (1987) Characterisation of the channel properties of a neuronal acetylcholine receptor reconstituted into planar lipid bilayers. J. Gen. Physiol. 90:855-879. Harrow, I.D., and D.B. Sattelle (1983) Acetylcholine receptors on the cell body membrane of giant interneurone 2 in the cockroach Periplaneta americana. J. Exp. Biol. 105:339-350. Hildebrand, J.G., L.M. Hall, and B.C. Osmond (1979) Distribution of binding sites for (1251)-labeleda-bungarotoxin in normal and deafferented antennal lobes ofManduca sezta. Proc. Natl. Acad. Sci. USA 76t499-503. Jacob, M.H., D.K. Berg, and J.M. Lindstrom (1984) Shared antigenic determinant between the Electrophorus acetylcholine receptor and a synaptic component on chick ciliary ganglion neurons. Proc. Natl. Acad. Sci. USA 81:3223-3227. Katz, B., and R. Miledi (1972) The statistical nature of the acetylcholine potential and its molecular components. J. Physiol. 224:665-699. Kehoe, J., R. Sealock, and C. Bon (1976) Effects of a-toxins from Bungarus multicinctus and Bungarus caeruleus on cholinergic responses in Apylsia neurons. Brain Res. 107:527-540. Kelly, J.S., and M.A. Rogawski (1985) I n M.A. Rogawski and J.L. Barker (eds): Neurotransmitter Actions in the Vertebrate Nervous System. New York Plenum Press, pp. 143-197. Kreissl, S., and G. Bicker (1989) Histochemistry of acetylcholinesterase and immunocytochemistry of an acetylcholine receptor-like antigen in the brain of the honeybee. J. Comp. Neurol. 6:71-84. Leake, L.D., and R.J. Walker (1980) Invertebrate Neuropharmacology. Glasgow: Blackie. Lieberman, E.M., J. Villegas, and G.M. Villegas (1981) The nature of the membrane potential of glial cells associated with the medial giant axon of the crayfish. Neuroscience 6:261-271.

57 Lutz, E.M., and N.M. Tyrer (1988) Immunohistochemical localization of serotonin and choline acetyltransferase in sensory neurones of the locust. J. Comp. Neurol. 267r335-342. Marks, M.J., and A.C. Collins (1982) Characterization of nicotinic binding in mouse brain and comparison with the binding of alpha-bungarotoxinand quinuclidinyl benzilate. Molec. Pharmacol. 22554-561. Marshall, L.M. (1981)Synaptic localization ofa-bungarotoxin bindingwhich blocks nicotinic transmission at frog sympathetic neurons. Proc. Natl. Acad. Sci. USA 78:1948-1952. Matthews-Bellinger, J., and M.M. Salpeter (1978) Distribution of acetylcholine receptors at frog neuromuscular junctions with a discussion of some physiological implications. J. Physiol. 279:197-213. Meyer, M.R., and G.R. Reddy (1985) Muscarinic and nicotinic cholinergic binding sites in the terminal ganglion of the cricket Acheta domesticus. J. Neurochem. 45t1101-1112. Morley, B.J., and G.E. Kemp (1981) Characterization of a putative nicotinic acetylcholine receptor i n mammalian brain. Brain Res. Revs. 3331-104. Murphy, S., and B. Pearce (1987) Functional receptors for neurotransmitters on astroglial cells. Neuroscience 22381-394. Oswald, R.E., and J.A. Freeman (1981) Alpha bungarotoxin binding and central nervous system nicotinic acetylcholine receptors. Neuroscience 6;l-14. Pearce, B. (1991) Glia as targets for neuroactive substances. In N.J. Abbott (ed): Glial-Neuronal Interaction. Ann. N.Y. Acad. Sci. 633:432433. Pellegrino, M., and M. Simonneau (1984) Distribution of receptors for acetylcholine and 5-hydroxytryptamine on identified leech neurones growing in culture. J. Physiol. 352669-684. Peper, K., and U.J. McMahon (1972) Distribution of acetylcholine receptors in the vicinity of nerve terminals on skeletal muscle of the frog. Proc. R. SOC.Lond. B. 181t431-440. Pfliiger, H.J., P. Braunig, and R. Hustert (1988) The organization of mechanosensory neuropiles in locust thoracic ganglia. Phil. Trans. Roy. SOC.B. 321:l-26. Rang, H.P. (1981) The characteristics of synaptic currents and responses to acetylcholine of rat submandibular ganglion cells. J. Physiol. 311.23-55. Rawlins, F.F.A., and J. Villegas (1978) Autoradiographic localization of acetylcholine receptors in the Schwann cell membrane of the squid nerve fibre. J. Cell. Biol. 77r371-376. Sattelle, D.B., and H. Breer (1990) Cholinergic nerve terminals in the central nervous system of insects. J. Neuroendocrinol. 2:241-256. Sattelle, D.B., J.A. David, I.D. Harrow, and B. Hue (1980) Actions of a-bungarotoxin on identified insect central neurons. In D.B. Sattelle, L.M. Hall, and J.G. Hildebrand (eds): Receptors for Neurotransmitter, Hormones, and Pheromones in Insects. Amsterdam: ElsevieriNorth Holland, pp. 125-139. Sattelle, D.B., I.D. Harrow, B. Hue, M. Pelhate, J.I. Gepner, and L.M. Hall (1983) a-Bungarotoxin blocks excitatory synaptic transmission between cercal sensory neurones and giant interneurone 2 of the cockroach, Periplaneta americana. J. Exp. Biol. 107t473-489. Sattelle, D.B., U. Madler, H. Heilgenberg, and H. Breer (1989)Immunocytochemical localization of nicotinic acetylcholine receptors in the terminal abdominal ganglion of the cockroach (Periplaneta americana). Proc. R. Sac. Lond. B. 238r189-192. Schmidt-Nielsen, B.K., J.I. Gepner, N.N.H. Teng, and L.M. Hall (1977) Characterisation of an a-bungarotoxinbinding component from Drosophila melanogaster. J. Neurochem. 29:1013-1029. Schuetze, S.M., and L.W. Role (1987) Developmental regulation of nicotinic acetylcholine receptors. Ann. Rev. Neurosci. 10:403457. Sombati, S., and G. Hoyle (1984) Glutamatergic central nervous transmission in locusts. J. Neurobiol. 15:507-516. Sternberger, L.A. (1974) Immunocytochemistry. New Jersey: Prentice Hall Inc. Strausfeld, N.J. (1976)Atlas of an Insect Brain. Berlin: Springer-Verlag. Tareilus, E., W. Hanke, and H. Breer (1990) Neuronal acetylcholine receptor channels from insects: A comparative electrophysiological study. J. Comp. Physiol. A 167:521-526. Tyrer, N.M., and G.E. Gregory (1982) A guide to the neuroanatomy of locust suboesophageal and thoracic ganglia. Phil. Trans. R. SOC.Lond. B. 297:91-123. Tzartos, S., and J.M. Lindstrom (1980) Monoclonal antibodies used to probe acetylcholine receptor structure: Localization of the main immunogenic region and detection of similarities between subunits. Proc. Natl. Acad. Sci. USA 77:755-759. Tzartos, S., L. Langeberg, S. Hochschwender, and J.M. Lindstrom (1983)

58 Demonstration of a main immunogenic region on acetylcholine receptors from human muscle using monoclonal antibodies to human receptor. FEBS Lett. 158:116118. Tzartos, S., D.E. Rand, B.L. Einarson, and J.M. Lindstrom (1981)Mapping of surface structures of Electrophorus acetylcholine receptor using monoclonal antibodies. J.Biol. Chem. 256:8635-8645. Usherwood, P.N.R. (1980) Neuromuscular transmitter receptors of insect muscle. In D.B. Sattelle, L.M. Hall, and J.G. Hildebrand (eds): Receptors for Neurotransmitter, Hormones, and Pheromones in Insects. Amsterdam: ElsevieriNorth Holland, pp. 141-160. Van Calker, D., and B. Hamprecht (1980) Effects of neurohormones on glial cells. Adv. Cell. Neurohiol. 1~31-67.

B. LEITCH ET AL. Villegas, J. (1984) Axon-Schwann cell relationship. Curr. Top. Membr. Transport. 2 2 5 4 7 6 7 1 . Villegas, J., and D.J. Jenden (1979) Acetylcholine content of the Schwann cell and axon in the giant nerve fibre of the squid. J. Neurochem. 32761-766. Wafford, K.A., D. Bai, M.I. Sepulveda, and D.B. Sattelle (1991) L-Glutamate receptors in the insect central nervous system. In B.S. Meldrum, F. Moroni, R.P. Simon, and J.H. Woods (eds): Excitatory Amino Acids. New York: Raven Press Ltd., pp. 275-279. Zatz, M., and M.J. Brownstein (1981)Injection of a-bungarotoxin near the superchiasmiatic nucleus blocks the effects of light on nocturnal pineal enzyme activity. Brain Res. 213:438-442.