No title

international Review of

NEUROBIOLOGY VOLUME 33

Editorial Board W. Ross ADEY

PAULJANSSEN

JULIUSAXELROD

KETY SEYMOUR

Ross BALDESSAKINI

KEITH KILLAM

SIRROGERBANNISTER

CONANKORNETSKY

FLOYDBLOOM

ABELLAJTHA

DANIELBOVET

BORISLEBEDEV

PHILLIP BKADLEY

PAULMANDEL

YURI BUROV

HUMPHRY OSMOND

Josf DELGADO

RODOLFOPAOLETTI

SIRJOHN ECCLES

SOLOMON SNYDER

JOELELKES

STEPHENSZARA

H . J. EYSENCK

MARATVARTANIAN

KJELLFUXE

STEPHENWAXMAN

Bo HOLMSTEDT

RICHARDWYATT

International Review of

NEUROBIOLOGY Editedby JOHN R. SMYTHIES Department of Neuropsychiatry Institute of Neurology National Hospital London, England

RONALD J. BRADLEY Department of Psychiatry and Behavioral Neurobiology The Medical Center University of Alabama Birmingham, Alabama

VOLUME 33

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9 8 1 6 5 4 3 2 1

CONTENTS

Olfaction

S. G. SHIRLEY I. Introduction . . . . . . . . . . . . . . . . ........................ 11. Biochemistry of Transduction ........................ 111. Physiology of Receptor Cells.. .................................. IV. Receptors and Patterns of Response.. ............................. V. Transfer of Information ......................................... VI. Perireceptor Events. ................ .......................... VII. Conclusion.. .................................................... References. . . . ......................................

1

7 13 16 25 35 40 41

Neuropharmacologic and Behavioral Actions of Clonidine: Interactions with Central Neurotransmitters

JERRYJ. BUCCAFUSCO I. 11. 111. IV. V. V1. VII.

Introduction .................................................... Receptor Specificity.. ........................... ........... Role of Brain Neurotransmitters in the Antihyperte Antiwithdrawal Effects. ......................... Other Pharmacological Actions ........................ Summary and Conclusions ....................................... Future Directions.. .. ......................... References . . . . . . . . . . . . . . . . . . . ..............................

56 59 63 73 85 95 98 100

Development of the Leech Nervous System

GUNTHERS. STENT,WILLIAMB. KRISTAN, JR., STEVENA. TORRENCE, KATHLEEN A. FRENCH, AND DAVIDA. WEISBLAT I. Introduction to the Leech.. ...................................... 109 11. Morphological Development and Staging .......................... 127 V

vi

CONTENTS

111. Behavioral Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Developmental Cell Lineage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Myogenesis and Neurogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I\’.

I34 137 151

183 187

GABA, Receptors Control the Excitability of Neuronal Populations

ARMIKSTELZER I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I95

11. GABAergic Inhibition: 111. GABAergic inhibition:

197 202

GABAA Receptor Function: Control of the Excitability o Populations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. GABA,\ Receptor Function: Tetanization .......................... VI. GABA;\ Receptor Function: Synchronization. ...................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

207 218 263 278

I\’.

Cellular and Molecular Physiology of Alcohol Actions in the Nervous System

FORREST F. WEIGHT ................................................ ............................... 111. Alcohol Effects on Neuronal Firing . . I\‘. Alcohol Effects on Cellular Mechanisms ........................... V. Summary and Conclusions . . . . . . . . . . ............ References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . CONTENTS OF RECENTVOLUMES ...........................

289 290 292 303 336 342

349 365

0LFACTl0 N S.

G.Shirley

Department of Chemistry University of Warwick Coventry, CV47AL, England

I. Introduction 11.

111.

IV.

V.

VI.

VII.

A. Scope B. Structure of the Peripheral Olfactory System Biochemistry of Transduction A. Cyclic AMP B. Phosphoinositides C. Direct Gating D. Lipid Responses Physiology of Receptor Cells A. Non-Odor-Induced Activity B. Odor- and Cyclic Nucleotide-Induced Activity Receptors and Patterns of Response A. Receptor Molecules B. Receptor Cells C. Responses in the Olfactory Bulb Transfer of Information A. The Vectorial Representation B. Odor Concentration and Signal/Noise C. Information Transfer from Mucus to Receptor Molecule D. Information Transfer from Receptor Molecule to Cell Perireceptor Events A. The Olfactory Mucus B. Control of Secretion C. Central Control of the Sensory Cells D. Xenobiotic-Metabolizing Enzymes E. Odorant-Binding Protein Conclusion References

1. Introduction

A. SCOPE The last decade has seen much progress in the understanding of olfaction. The purpose of this article is to review the molecular and cellular processes by which information is extracted from an incoming I INTERNATIONAL REVIEW OF NEUROBIOIDGY, VOL. 33

Copyright 0 1992 by Academic Press, lnc. All rights of reproduction in any form reserved.

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S. C . SHIRLEY

olfactory stimulus and passed to the central nervous system. T h e mechanisms of insect olfaction are excluded and readers are directed to the works of Kaissling (1986, 1987), O’Connel (1986), and Payne et al. (1986). Although work on invertebrates and fish is mentioned, this article concentrates mainly on the processes occurring in the air-breathing vertebrates.

€3. STRUCTURE OF THE PERIPHERAL OLFACTORY SYSTEM

1 . Oueroiew I n mammals and amphibia, the sensory cells of olfaction lie within the olfactory mucosa. In mammals, the mucosa covers part of the ethnioturbinates and part of the nasal septum, so forming the lining of airfilled cavities adjacent to the main air passages. During inhalation, eddies break away from the main airstream and enter these cavities, where airborne organic molecules can interact with the olfactory receptors. T h e superficial layer of the mucosa is a pseudostratified columnar epithelium. It is avascular and contains the sensory cells, supporting cells, and basal cells. A thin layer of mucus covers the olfactory epithelium, separating it from the air space. T h e mucus derives from the supporting cells and olfactory glands. T h e deeper part of the mucosa, the lamina propria, contains blood vessels, connective tissue, secretory glands, and bundles of sensory cell axons. The ultrastructure of the olfactory mucosa has been reviewed by Menco (1983). Morphologically and functionally the sensory cells are bipolar neurons, although they are, embryologically, of epithelial origin (Cuschieri and Bannister, 1975). Axons from these cells run in bundles or fasciculi through fine holes in the cribiform plate and into the brain. T h e axons form synapses within the glomeruli of the olfactory bulb. T h e glomeruli are large structures (loo+ pm) where axons from several tens of thousands of sensory cells form en passant synapses with about 100 secondary (mitral and tufted) cells. T h e neurotransmitter at these synapses is the dipeptide carnosine (P-alanyl- 1-histidine) (Neidle and Kandera, 1974; Gonzales-Estrada and Freeman, 1980; Burd et al., 1982; Margolis et al., 1985, 1987; Sakai et al., 1987, 1988). The axons of the mitral cells and of some tufted cells are the main output of the olfactory bulb and run ipsilaterally to the olfactory cortex. In addition to the olfactory system outlined above, most air-breathing animals (excluding man) have an additional chemical sense, the vomeronasal system. This usually takes the form of a tube, lined with

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3

sensory cells, linking the oral and nasal cavities. The axons of these cells project to the accessory olfactory bulb; this structure is generally similar to and contiguous with the olfactory bulb. Whereas the vomeronasal system is usually stimulated by materials in solution, it can be stimulated by airborne substances, probably after solution in saliva. The importance of this system has been discussed by Wysocki (1979) and Doty (1988). There is also innervation of the nasal cavities by the trigeminal nerve, and it plays a role in chemosensation (e.g., Silver et al., 1986). 2. The Primary Cells The sensory cell bodies are fusiform, small (typically 15 X 4 pm), and generally lie at depths of greater than about 50 pm within the olfactory epithelium. From each, a single dendrite extends between the supporting cell to the tissue surface and ends in a terminal swelling about 1.5 pm in diameter. The terminal swellings carry modified cilia. T h e basal bodies of these cilia appear normal in the electron microscope. Each cilium has a proximal segment about 1.5 pm in length and 200 nm in diameter containing the usual 9(2) + 2 pattern of filaments. The cilia then taper to about 90 nm in diameter and the filaments become fewer in number and less regular in organization. In mammals, the distal parts of the cilia are typically 30 pm in length; in amphibia they may be up to 100 pm long. Mature olfactory cilia are immotile, although immature forms of amphibian cilia may be motile (Mair et al., 1982). Scanning electron microscopy has proved valuable in revealing details of the olfactory surface in fish (Breipohl et al., 1973; Doroshenko and Motavkin, 1986), amphibians (Mair et al., 1982; Menco, 1977; Klein and Graziadei, 1983), birds (Breipohl and Fernandez, 1977), reptiles (Wang and Halpern, 1980), and mammals (Menco, 1977). The technique can also be used to reveal the structure of the mucosa in depth (Costanzo and Morrison, 1989; Morrison and Costanzo, 1989). Measurements of the period between odor application and response, coupled with a knowledge of the diffusion coefficients of the odorant, imply that the transductory apparatus is very close to the tissue surface, in the cilia, terminal swellings, or apical part of the dendrite (Getchell et al., 1980). Other evidence reviewed by Getchell et al. (1984) also suggests that the transductory apparatus is located in the cilia. At the level of the ciliary taper there is a prominent necklace of particles in the membrane. The necklace particles are firmly anchored to the cytoskeleton. They form a region of membrane attachment and a barrier to lateral diffusion in the membrane (Menco, 1988b). T h e distal parts of the cilium have a high density of membrane-embedded particles (Menco et al., 1976). Particle densities of 1000 pm-* (frog) to 2300

4

S . G. SHIRLEY

pm-2 (dog) were found in a series of freeze-fracture studies by Menco (198Oa-d). It is hypothesized that these particles form part of the transductory mechanism. T h e cilia1 membranes of the receptor cells are also more electron opaque than those of the respiratory cells (Menco, 1989b). Freeze-fracture techniques have also been used to study the monkey epithelium by Engstriim et al. (1989). The olfactory receptor neurons are unusual in that they have a finite lifetime and they are replaced by new cells derived from the basal cells. Breipohl et al. (1986) have reviewed the control mechanisms of this replacement. I t is possible that the availability of space for the developing neuron is a controlling factor (Mackdy-Sim et al., 1988). It is likely under natural circumstances that each receptor cell has a lifetime of many months (Hinds et al., 1984), but this can be artificially reduced. Following transection of the axons, the receptor cells degenerate. New cells appear at about the tenth day and replacement is complete at about a month. Simmons and Getchell(l981) found that the newly differentiated neurons had spontaneous electrical activity, odor specificity, and concentration-response profiles as normal neurons before the axons made contact with the olfactory bulb. However, Masukawa et al. (1985) found that the newly differentiated cells were generally nonspiking. Immature cells have been found to be less responsive to odor than are mature ones and they show few action potentials (Lidow et al., 1987a,b); they also have smaller resting potentials and are less responsive to injected current (Hedlund et al., 1987; Masukawa et al., 1987). The axons of the sensory cells are nonmyelinated; Schwann cells envelop bundles of axons and extend tongues of cytoplasm between the fibers (Barber and Lindsay, 1982; Rafols and Getchell, 1983). There are conflicting reports as to whether the Schwann cells are epithelial or glial. Vollrath et al. (1985), Ophir (1987), and Ophir and Lancet (1988) found keratin as the filament protein. Whereas glial fibrillary acidic protein (GFAP) was found by Barber and Lindsay (1982) and Barber and Dahl (1987), who reported that there were low levels of the central type of GFAP for much of the length of the fasciculi but higher levels at the receptor cell end and the possibility that peripheral-type GFAP is also present. I n view of the continuous turnover of the olfactory neurons, several groups have looked for signs of immaturity in these cells. T h e cells have immature versions of microtubule-associated proteins (Viereck et al., 1989) and cell adhesion molecules (Miragall et al., 1988). It is rare for olfactory axons to show staining with antibodies to neurofilament protein (Vollrath et al., 1985), but there is disagreement as to the presence of vimentin. This intermediate filament protein has been reported by

OLFACTION

5

Schwob et al. (1986) but not by Vollrath et al. (1985), Ophir (1987), or Ophir and Lancet (1988) (see also Yamagishi et al., 1989a,b). Antigens associated with neuritic outgrowth continue to be expressed in the adult animal (Stallcup et al., 1985; Wallis et al., 1985). The growth-associated protein B50/GAP43 can be found in the epithelium of adult rats (Verhaagen et al., 1989). It seems to be associated with the newly differentiated subpopulation of cells and its expression ceases when that of olfactory marker protein (OMP) starts. OMP is a protein of unknown function, taken to be a marker of mature olfactory receptor neurons (Margolis, 1972, 1988), but is also found at very much lower levels in other brain regions (Baker et al., 1989). The expression of OMP is thought to coincide with ciliogenesis (Menco, 1989a). The olfactory receptor neurons also carry receptors for nerve growth factor (Taniuchi et al., 1986) and high levels of nerve growth factor can be found within the neurons of the developing epithelium (Williams and Rush, 1988). Regenerative neurogenesis is not a recapitulation of prenatal development (Menco, 1985). Development processes have been reviewed by Brunjes and Frazier (1986), Penderson et al. (1986a), Farbman and Menco (1986), Farbman (1988), and Mair (1988). Regeneration following transplant or bulbectomy has been reviewed by Graziadei and MontiGraziadei (1986a) and Barber and Jensen (1988). It is likely that under normal circumstances the axons reach their targets along preexisting glial channels (Barber, 1982a,b; Barber and Dahl, 1987; MontiGraziadei and Morrison, 1988; Monti-Graziadei and Graziadei, 1989). But it has been hypothesized by Graziadei and Monti-Graziadei (1986b) that no target is necessary for glomerulus formation, the receptor cell axons being capable of self-selection. In addition to the ciliated cells already described, there are also some microvilli-bearing cells with distal processes (e.g., Menco, 1980a; Moran et al., 1982; see also Akerson, 1988) and it has been demonstrated by Rowley et al. (1989) that these send axons to the olfactory bulb. The function of these cells is unknown, but in the human 10% of axons from the olfactory region derive from such cells.

3. Connections between the Epithelium and Bulb There is a relationship between the position of a glomerulus and the positions of its sensory cells. To each glomerulus corresponds a field that is a set of strips running anterior-posterior in the epithelium. By no means do all sensory cells in such a strip project to the same glomerulus, but there is a tendency for them to project to neighboring ones. This has been demonstrated by horseradish peroxidase injection or lavage Uastreboff et al., 1984; Stewart, 1985; Penderson et al., 1986b; Astic and

6

S. G . SHIRLEY

Saucier, 1986, 1988; Saucier and Astic, 1986; Astic et al., 1987; Stewart and Penderson, 1987; Zheng and Jourdan, 1988), radio-leucine injection (Mackay-Sim and Nathan, 1984), and physiological methods (Costanzo and Mozell, 1976; Costanzo and O’Connel, 1980). Any particular bulb neuron has a field of receptor cells that are excitatory and a field of cells that are suppressive and there is a tendency for the excitatory field to be more compact (Kauer and Moulton, 1974). There is electrophysiologicdl evidence that the excitatory and inhibitory components of the receptive field of an olfactory bulb neuron are not well segregated Uiang and Holley, 1987). I n addition to dense projections to neighboring glomeruli, any given region of epithelium has diffuse projections to the whole olfactory bulb (Kauer, 1981) and there are diffuse projections from the whole epithelium to any part of the bulb (Kauer, 1980). There are monoclonal antibodies capable of distinguishing subclasses of olfactory receptor cells and there is a tendency for any given glomerulus to receive its input from like cells (Fujita et al., 1985; Mori et al., 1985; Mori, 1987b; Schwob and Gottlieb, 1986, 1987, 1988) o r for there to be a general relationship between the position of the positive cells and the positive glomeruli (Shinoda et al., 1989). T h e same kind of relationship may be observed with lectin binding (Key and Giorgi, 1986; Barber, 1989). 4. The Olfclctoq Bulb

The structure of the olfactory bulb has been reviewed by Halasz and Shepherd (1983), Macrides and Davis (1983), Scott (1986), Mori (1987a), Scott and Harrison (1987), and Dawson et al. (1988). T h e principal cells are the mitral and the tufted cells. Each extends a single primary dendrite that arborizes within a single glomerulus and receives input from the sensory cell axons. Each side of the rabbit olfactory bulb has about 1900 glomeruli with about 24 mitral and 70 tufted cells per glomerulus. Each glomerulus receives the axons of about 25,000 sensory cells and the en p a m n l synapses are such that each mitral cell receives input from 1300-6500 sensory cells. This extremely high degree of convergence is one of the characteristic features of the olfactory system. In addition, each mitral or tufted cell extends secondary dendrites that make many reciprocal dendrodendritic synapses with the axonless granule cells. The olfactory bulb is very rich in dendrodendritic synapses and is a useful source of material for their study (Chiflikian et al., 1986). I‘he granule-to-mitral synapse is inhibitory, using y-aniinobutyric acid as transmitter. T h e niitral-to-granule transmission, which includes some axodendritic synapses, is excitatory. T h e main axons of the mitral and

OLFACTION

7

some tufted cells project to the ipsilateral olfactory cortex. Collaterals branch, forming synapses with granule cells. These collaterals extend further than do the secondary dendrites of the same cell and some project to the other side of the olfactory bulb, making synapses with granule cells. There are subtypes of mitral and tufted cells distinguishable on morphological and functional grounds (Macrides et al., 1985; Scott, 1986, 1987). Within the glomeruli; the sensory cell axons also form synapses with the dendrites of periglomerular cells. There are indications that the sensory cells induce a dopaminergic phenotype in the periglomerular cells via a calcitonin gene-related peptide (Denis-Donini, 1989; see also Baker, 1988). T h e periglomerular cells also form reciprocal dendrodendritic synapses with the mitral and tufted cells. The periglomerular cells have short axons, which make synapses with the primary dendrites of the mitral and tufted cells and with the dendrites and somata of other periglomerular cells. There are also several other types of short axon cells within the olfactory bulb. The main centrifugal input to the olfactory bulb is to the granule cells. Many neuropeptides, hormones, or their receptors have been identified in the olfactory bulb. These include insulin (Matsumoto and Rhoads, 1990; see also Baskin et al., 1988), atrial natriuretic peptide (Katawa et al., 1986; Gibson et al., 1988; Glembotski et al., 1989), neuropeptide Y (Scott et al., 1987; Ohm et al., 1988), somatostatin (Scott et al., 1987), enkephalins (Kosaka et al., 1987), substance P (Baker, 1986a,b), vasoactive intestinal polypeptide (Alonso et al., 1989), luteinizing hormone-releasing hormone (Zheng et al., 1988), and cholecystokinin-8 (Seroogy et al., 1985; Matsutani et al., 1989a). The distributions of many of these substances define subpopulations of cells in the bulb and the distributions of several neuropeptides have been studied in the guinea pig (Matsutani et al., 1989b) and in the developing rat (Matsutani et al., 1988).

11. Biochemistry of Transduction

This subject has been reviewed (Lancet, 1986, 1988; Lancet et al., 1988; Bruch et al., 1988; Snyder et al., 1988b; Shirley and Persaud, 1990) and there have been several minireviews (Anholt, 1987; Lancet and Pace, 1987; Snyder et al., 1988a, 1989). Four kinds of transduction mechanisms have been proposed. These are mechanisms based on cyclic

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S. G . SHIRLEY

AMP, phosphoinositides, the direct opening of ion gates by odorants, and the inherent response of lipids to odorants. These will be discussed in turn.

AMP A. CYCLIC In general, adenylate cyclase systems consist of a number of components: one or more types of receptor, one or more guanine nucleotidebinding proteins (G proteins), the adenylate cyclase catalytic unit, and one o r more phosphodiesterases. Normally the G protein carries tightly bound guanosine diphosphate (GDP). T h e binding of a ligand to a receptor allows the G protein to exchange this for cytosolic guanosine triphosphate (GTP). If the G protein is of the stimulatory type (G,), this activated form of the G protein interacts with the catalytic unit to increase catalytic activity. T h e G protein hydrolyzes its bound GTP to GDP and returns to the inactive form. Inhibitory G proteins (G,) are believed to act by deactivating G , (e.g., Gilman, 1987), but to date there is no evidence for the involvement of an inhibitory G protein in olfactory transduction. The adenylate cyclase catalytic unit converts ATP to cyclic AMP; this is the only known mechanism for the generation of this species. Cyclic AMP has a controlling function for many cellular processes and it is converted to AMP by the action of the phosphodiesterases. In addition to its transductory function, cyclic AMP has been implicated in the control of neuritic outgrowth in developing cells. An analog of cyclic AMP, phosphodiesterase inhibitors, and forskolin (a potent stimulant of adenylate cyclase) all promote outgrowth in explanted developing olfactory epithelia (Johnson et al., 1988b). 1. E v d e i t c e f o r the Inzdvement of A d e y late Cyclase

Kurihara and Koyania (1972) demonstrated the presence of high levels of adenylate cyclase in the olfactory mucosa. The olfactory cilia of mammals and amphibia can be detached by calcium shock (Anholt et al., 1986; Chen et al., 1986a; Lazard et al., 1989) o r by sonication (Shirley et al., 1986). Such preparations show a very high level of adenylate cyclase activity, which can be stimulated by the addition of odorants (Pace et al., 1985; Shirley et al., 1986; Sklar et al., 1986). The dose-response curves for odorant activation of adenylate cyclase are fairly flat, having Hill coefficients of about 0.3-0.6, and so are similar in shape to overall olfactory dose-response curves from psychophysical measurements. The odorant concentration range in which stimulation of the adenyl-

OLFACTION

9

ate cyclase occurs is typically micro- to millimolar. It is important to remember that despite their generally hydrophobic nature, most odorants partition so that their concentration in water (or mucus) is much higher than that in air (see Section V1,A). This concentration range is comparable with that which will elicit an electrophysiological response from the tissue (Shirley et al., 1987b) and, when corrected for partitioning, that which can be smelled. There is a good correlation between an odorant’s potency in stimulating adenylate cyclase and its potency in stimulating the electroolfactogram (Lowe et al., 1989). The electroolfactogram is a measure of the combined generator currents of the receptor cells (Ottoson, 1956; Gesteland, 1975). The report by Sklar et al. (1987) that some classes of odorant do not stimulate adenylate cyclase is based on measurements at a single odor concentration, which may have been subthreshold for some compounds. Some materials classified as nonstimulants by Sklar et al. (1987) have been found to stimulate at higher concentrations (Lowe et al., 1989). Adenylate cyclase has been histochemically localized to the cilia, terminal swellings, and dendritic shafts of the primary cells (Asanuma and Nomura, 1989). Jones et al. (1988) have shown that both the enzymatic activity and the mRNA for adenylate cyclase are predominantly expressed in the receptor cells. Wheat germ agglutinin, which inhibits the electrophysiological response to odorants, also inhibits the odorant activation of adenylate cyclase (Lancet et al., 1987). Cyclic AMP analogs and phosphodiesterase inhibitors have been shown to interfere with olfactory transduction by Minor and Sakina (1973), Menevse et al. (1977), and Persaud et al. (1988a). There is good evidence (Sections III,B,2 and III,B,3) for the existence of cyclic nucleotide-controlled ion gates in the sensory cells. There have been no reports of measurement of cyclic AMP levels within the olfactory tissue, but neuroblasts from the olfactory epithelium have been shown to accumulate cyclic AMP when challenged with odorants in culture (Coon et al., 1989). Preparations of olfactory cilia also contain proteins that are phosphorylable in a cyclic nucleotide-dependent manner (Heldman and Lancet, 1986; Kropf et al., 1987). T h e adenylate cyclase catalytic unit of the olfactory system has been isolated by Pfeuffer et al. (1989). It is a novel form of the enzyme, having a high molecular weight (180,000), and it is highly active, with a turnover number of about 8000Imin.

2. G Protein Guanosine triphosphate is essential for the activation of the olfactory adenylate cyclase (Pace et al., 1985; Shirley et al., 1986), implying that the

10

S. G. SHIRLEY

cyclase is coupled to the receptor via a G protein. This is confirmed by the observation of Vodyanoy and Vodyanoy (1987a,b) that lipid bilayers incorporating olfactory membrane fragments require GTP in order to display conductance changes; Weinstock et al. (1986) and Wright et al. (1987) also confirmed that humans with a G protein deficiency have an impaired sense of smell. Further evidence for the involvement of a G protein in transduction comes from the observation that odorants can cause small modulations in the binding of guanine nucleotide analogs in olfactory preparations (Lancet, 1988) and by guanine nucleotide-induced changes in the affinity of olfactory receptors for ligdnds (Bruch and Kalinoski, 1987). NotJoselov et al. (l988a,b) have isolated a protein complex from the skate. T h e complex can be dissociated by GTP or analogs and its GTPase activity can be increased by amino acids that are olfactory stimuli for this fish. This could be a receptor-(; protein complex. Interaction between G protein and the cyclase catalytic unit can be demonstrated in reconstituted detergent extracts of olfactory material (Anholt, 1988). The involvement of G proteins in olfaction has also been studied by Parfenova and Etinghof (1988). Several G proteins can be found in the olfactory mucosa. T h e stimulatory (GJ, inhibitory (G,), and other (Go) varieties are present and G, is enriched in cilia1 preparations (Anholt et al., 1987a,b). However, the G protein involved in olfactory transduction is a variant (Pace and Lancet, 1986). It has now been cloned and is called G<,,, (Jones and Reed, 1987, 1989).There is evidence that Gp, the subunit common to all G proteins, and its mRNA are present in high levels in the olfactory receptor cells (Jones et al., 1988) and that Gp is found in very high levels at the tissue surface (Anholt at al., 1987a,b). Pxe and Lancet (1986) point out that olfactory preparations are unusual among preparations of neural cyclases in that a stimulatory G protein is the predominant species. Another unusual property is the very high apparent affinity shown by the G protein for guanine nucleotides (Pace and Lancet, 1986; Anholt et al., 1987b; Robinson et al., 1989). This may be a system property rather than a molecular one and arises because of a large stoichiometric excess of G protein over cyclase catalytic units (Robinson et al., 1989). Further, this excess is partially due to the fact that 85-90cJc of the cyclase is normally decoupled from the C; protein and so remains silent. If coupling is artificially induced, the apparent affinity drops toward more normal values. It is not known if this is of physiological significance, but it could indicate the presence of a sensitivity control located very close to the periphery of the system.

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11

3. Phosphodiesterases Little work has so far been done on the phosphodiesterases of the olfactory tissue. Both high- and low-affinity isoenzymes are present; they are not odor sensitive and at least one isoenzyme is stimulated by calcium via calmodulin (Dickinson, 1987).A low-affinity form of the enzyme may be stimulated by estradiol (Parfenova, 1987). 4. Calcium

It is likely that calcium plays an important part in the transductory process (e.g., Suzuki, 1978). There are calcium currents in the primary cells (Section III,A,l), and reports show that calcium plays an important role in the process of adaptation (Kurahashi and Shibua, 1989; Kurahashi, 1989; Suzuki, 1989). This may be linked to the observation that calcium is inhibitory to adenylate cyclase (Shirley et al., 1986; Sklar et al., 1986) and stimulatory to phosphodiesterase (Dickinson, 1987). Odorant stimulation of the cyclase and the increased cAMP levels eventually cause depolarization of the cell, and it is possible that this is accompanied by increased calcium levels, which by inhibition of the cyclase and stimulation of the phosphodiesterase reduce the cAMP concentration. The calcium ion, by playing the role of a feedback regulator (with lag), would impart to the system some degree of “edge enhancement,” rendering it more sensitive to increasing odor concentrations than to steady ones. This is in addition to the effect of calcium on the ion channels of the cell (Section III,A,l). I n view of the possible importance of calcium, it is interesting that S-lOOpp, a member of the calcium regulatory protein family and normally found only in glial cells, is present in these neurons (Rambotti et al., 1989). There is also a report that high levels of calcium and calcium channel blockers can inhibit transduction at very high odorant levels (Winegar et al., 1988).

B. PHOSPHOINOSITIDES Phosphoinositides are important second messengers in many cells (reviewed by Hokin, 1985). There are preliminary reports that phosphoinositides may be important in olfactory transduction in waterbreathing animals. Phospholipase C is present in the olfactory cilia of the catfish (Boyle et al., 1987) and alanine (an olfactory stimulus for these animals) can stimulate phosphoinositide turnover (Huque and Bruch, 1986; Bruch and Huque, 1987).The turnover is also stimulated by GTP,

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implying that a G protein may be involved, a conclusion reinforced by changes of ligand affinity in the presence of GTP (Bruch and Kalinoski, 1987). There are no rep0rt.s of phosphoinositide-based transduction mechanisms among the air-breathing animals.

C. DIRECTGATING Vodyanoy and Murphy ( 1983)discovered that homogenates of olfactory mucosa could impart an odor sensitivity to artificial bilayers. The active component was a 62-pS potassium channel that was directly activated by odorants at concentrations as low as 25 nM. The possibility of the direct action o f odorants on ion gates has also been discussed by Price (1984).Several ion channels have been found in bilayers fused with an olfactory cilia1 preparation (Labarch and Bacigalupo, 1988; Labarch et al., 1988). Among these was one that could be activated by odorants at concentrations down to 40 nM. The channel is described as a multistate cation channel, its conductance being 35 pS and multiples thereof.

D. LIPIDRESPONSES Kashiwayanagi et u1. (1987a,b) have shown that cells from the olfactory epithelium respond to odorants with changes of membrane fluidity and changes of membrane potential that are not dependent on sodium or chloride. Carp olfactory cells can show responses to odorants after removal of their cilia (Kashiwayanagi et ul., 1988), although deciliation of the frog olfactory epithelium has been shown to seriously impair function (Admek et al., 1984). Nonolfactory cells can display responses to odorants (Kashiwayanagi and Kurihara, 1984; Lerner el al., 1988) and biological membranes generally can respond in some fashion to small organic molecules (e.g., Schneider, 1968; Seeman et ul., 1971). Organic molecules in general partition into biological membranes, causing changes in membrane fluidity and hence changes in the activity of enzymes, including adenylate cyclase (e.g., Houslay and Gordon, 1983). Liposornes show membrane potential changes in the presence of odorants (Nomura and Kurihara, 1987a) and varying the lipid composition of the liposomes changes the response to different odorants differentially (Nomura and Kurihara, 1987b). There is ample evidence that lipid-mediated responses to odorants

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can occur and are not confined to the olfactory system. The question is whether they are an integral part of olfactory transduction, a complicating factor, o r an irrelevance. The proposed mechanism for the generation of the membrane potential change in bilayers is an alteration of the phase boundary potential of the membrane (Nomura and Kurihara, 1989). This is not a likely mechanism for transduction generally (Murphy, 1988) and does not accord with the physiological finding that there is a conductance change in the transducing membrane. But the ionic requirements for this mechanism are very nonrestrictive and it is conceivable that a freshwater fish might have evolved a transductory mechanism with this basis. The ionic requirements for olfaction in the carp (Kurihara and Yoshii, 1983) are similar to those of lipid bilayer responses. Price (1984) pointed out that whatever mechanism a cell used to transduce odor information, lipid-mediated effects would be superimposed upon its response. Such effects tend to increase sharply with concentration once a critical concentration is achieved and this could set an upper limit to the concentration range over which the olfactory system can function. T h e olfactory adenylate cyclase is unusually insensi.tive to changes of membrane fluidity (Shirley et al., 1987c), and this may be an adaptation to increase the system’s useful range. There is also some evidence that modification of cell surface proteins can change the response of the tissue differentially to different odorants (Section IV,A, l), implying that proteinaceous receptor sites are present. Whether o r not membrane fluidity modulations are important in olfactory transduction, the results of Nomura and Kurihara ( 198713) underline the theoretical point that an odor-discriminating system need not possess sharply tuned receptors (Section V,C,4). An analysis of the total lipid of the olfactory mucosa has been performed by Russell et al. (1989); the only unusual features are a slightly high proportion of polyunsaturated fatty acids and the presence of some transunsaturated fatty acids. But a lipid analysis on the cell-by-cell basis needed to understand a lipid-based transduction system is beyond current technology.

111. Physiology of Receptor Cells

The functional properties of the olfactory receptor neurons have been reviewed by Getchell (1986) and by Trotier (1990).

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A. NON-ODOR-INDUCED ACTIVITY

There have been several studies of the currents and ion gates necessary for the generation of action potentials in the olfactory cells. 1 . Membrane Currents

Firestein and Werblin (1987a), using a whole cell patch clamp on isolated receptor neurons, could identify five membrane currents. There was a transient inward sodium current that was insensitive to tetrodotoxin and a smaller sustained inward calcium current that was present at membrane potentials more positive than -20 mV. There were three outward potassium currents; one was calcium dependent, one was a slowly activating voltage-dependent current, and one was an inactivating potassium current. Firestein and Werblin (1987b) point out that this last current has characteristics that explain the tendency of the olfactory cells to produce spike trains of decreasing amplitude. The membrane resistance of these small cells is extremely high (3-6 GR) and currents of as little as 3 pA are sufficient to cause depolarization and action potential initiation. ‘These finding are similar to those of Trotier (1986) and similar results have also been obtained by Schild (1989), with the exception that tetrodotoxin blocks the transient inward sodium current, which may be a matter of species difference. Dionne (1987) and Suzuki (1987) also have broadly similar findings. Several authors have remarked on the very high sensitivity of the olfactory neurons. With their small size and high membrane resistance, these cells may be depolarized to the point of action potential initiation by the opening of a few ion channels or perhaps by the opening of a single ion channel (Firestein and Werblin, 1987b; Maue and Dionne, l987b; Kurahashi, 1989; Lynch and Barry, 1989; Schild, 1989).

2. Ion Gates Maue and Dionne (1987a,b, 1988) have identified several ion channels in mouse olfactory receptor neurons. There is a 130-pS potassium channel that is activated by calcium in the 0.5-1 ph4 range, an 80-pS potassium channel less sensitive to calcium found only in the adult mouse, and a voltage-activated potassium channel of 40 pS. In addition there is a 16-pS voltage-activated calcium channel and 2 10-pS chloride channel. T h e presence of a chloride channel in an invertebrate has been demonstrated by McClintock and Ache (1989a). With the reports by Yoshii and Kurihara (1989) and Getchell (1988) that Xenopus oocytes injected with niRNA from the olfactory epithelium can show an inward

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rectifier current and odor-induced current transients, respectively, there is promise that the power of molecular biological techniques will soon reveal details of the ion channels of olfactory cells.

B. ODORAND CYCLIC NUCLEOTIDE-INDUCED ACTIVITY 1. Whole Tissue Responses Using a whole tissue voltage-clamp technique, Persaud et al. (1987) found a transmucosal chloride current that was blocked by furosemide and was not odor sensitive. In addition, there were odor-induced currents. These were partially sensitive to amiloride and so probably were flowing through a cation channel with a degree of selectivity for sodium. T h e effect of cyclic AMP and other agents on the odor-induced current has been studied (Persaud et al., 1988a; DeSimone et al., 1988). Bromocyclic AMP, a membrane-penetrating cyclic AMP analog, when applied to the ciliated side of the mucosa (but not the submucosa), induced an inward current that was also partially blocked by amiloride, a drug that also blocks the generator current of isolated cells (Frings and Lindermann, 1988). T h e concentration of the analog necessary for half-maximal current was 2.75 p,M and the current-concentration curve had a Hill coefficient of 1.5. Bromo-cyclic AMP and odorant seem to activate the same current source and there was evidence for the involvement of a G protein in the generation of the odor-induced current. Interestingly, cyclic GMP was found to control an outward current.

2 . Whole Cell Responses Trotier (1986), using a patch clamp, showed that odor-induced currents passed through a nonselective cation channel and had a reversal potential of about 0 mV. A diffusable intermediate was probably involved in the transductory process. This confirms the finding of earlier work using intracellular recording techniques (Trotier and MacLeod, 1983) that the odor-gated current has a near-zero reversal potential and the finding of Getchell (1974) that both sodium and potassium are important as charge carriers (but see also Leveteau et al., 1989). Kurahashi (1989) has shown that isolated cells can gate currents of up to 400 pA when challenged with high concentrations of odorants. T h e dendrites and cilia seem to be the most odor-sensitive parts of the cell. The current flows through a nonselective monovalent cation channel and calcium is involved in the process of adaptation. There has been a demonstration that the odor-controlled current and spontaneous ac-

16

S. G. SHIRLEY

tivity of isolated cells can be blocked by amiloride (Frings and Lindermann, 1988), although the necessary concentration of the drug is very much higher than that needed to block other sodium currents. Maue and Dionne (1987b, 1988) described an odor-sensitive current in isolated mouse receptor cells. T h e current can also be stimulated by forskolin, a potent stimulator of adenylate cyclase. Such a response is not seen in excised patches and this negative result would be expected if a soluble species such as cyclic AMP were mediating transduction. An odor-induced generator current with near-zero reversal potential was observed by Anderson and Hamilton (1987) in isolated salamander cells, and such currents have been studied by Firestein and Werblin (1989) at odorant concentrations down to 6 ~J.M.Similar results have been obtained by Anderson and Ache (1985) and by Schmiedel-Jacob et al. (1 989) from the olfactory cells of the lobster. Trotier and MacLeod (1986a, 1987) have shown that forskolin and cyclic AMP can cause inward currents of several hundred picoamps in patch-clamped isolated salamander receptor neurons. Cyclic nucleotide-gated currents have also been observed by Vodyanoy (1988) and Suzuki (1988).

3. Ion Channels There is a report (Nakamura and Gold, 1987) that the olfactory cells contain an ion gate that passes a current with a reversal potential near 0 mV and opens by the direct binding of cyclic nucleotides, with no requirement for phosphorylation, and that this may be related to an ion gate of the visual system (Gold and Nakamura, 1987).

IV. Receptors and Patiems of Response

The striking characteristics of the olfactory system are its ability to respond to virtually any chemical species that may be delivered to it and its ability to discriminate between stimuli. This is best considered in terms of response spectra that describe the range of compounds to which any one cell or type of receptor molecule will respond and response patterns that describe the activity induced in many different cells or molecules by an odorant. Also important is the degree to which the response spectra of different cells or types of receptor molecules overlap. A cell or receptor molecule responsive to only a very limited range of stimuli is described as tightly tuned and so would be analogous to a receptor for a neurotransmitter or hormone. One stimulated by a large number of compounds is described as broadly tuned.

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A. RECEPTORMOLECULES

1. Response Spectra In fish, at least some of the receptor molecules exhibit fairly tight tuning. There are reports of correlations between amino acid binding and electrophysiological responses in catfish (Cancalon, 1978) and trout (Cagan and Zeiger, 1978; Brown and Hara, 1981, 1982). In the salmon there is a correlation between binding and behavior, which suggests that although some amino acids may share binding sites, the overlap is not extensive (Rehnberg and Schreck, 1986). The catfish possesses a receptor molecule responsive to L-alanine. Binding and competition experiments (Rulli and Bruch, 1987; Bruch and Rulli, 1988) have established that the receptor will accept other amino acids, preferring short-chain neutral molecules; it will, for instance, accept D-alanine at fourfold reduced affinity. These biochemical data parallel electrophysiologicaldata from the receptor cells, implying that these cells carry only one type of receptor molecule (or at least only one type that can respond to this kind of material). Tight tuning of receptor molecules has not been found in the airbreathing animals. The olfactory tissue can be treated with certain reagents that will modify its electrophysiological response to odorants. If such a reagent inactivated a tightly tuned receptor, the response to only a small group of closely related odorants should be affected. This perhaps occurs in the catfish (Kalinoski et al., 1987). In mammals, however, the pattern of experimental results is that responses to many odorants are affected to different degrees. This has been observed for membranepenetrant thiol reagents (Getchell and Gesteland, 1972; Delaleu and Holley, 1980), nonpenetrant thiol reagents (Menevseet al., 1978; Shirley et al., 1983), photoreactive odorants (Delaleu and Holley, 1983), enzymatic iodination (Shirley et al., 1983), lectins (Shirley et al., 1987a,b; Polak et al., 1989), and antibodies (Price and Willey, 1987). Price and Willey (1988) have shown that some antibodies, raised to candidate receptor molecules, are capable of reducing the electrophysiological response of the tissue selectively toward certain odorants and thus they may be supposed to recognize odor-binding sites of receptor molecules. But in these cases about two-thirds of the original response remains, implying a significant degree of overlap in response spectra. Other antibodies can abolish response to all odorants and may be recognizing general features of receptor molecules. The ability of some antibodies raised against proteins and other protein-specific reagents (Shirley et al., 1983) to modify responses in an odor-dependent

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SHIRLEY

fashion strongly implies the existence of proteinaceous receptor molecules. In some experiments, a large enough number of odorants was used to detect a pattern in the degrees of reduced response that may relate to response spectra (Delaleu and Holley, 1983; Shirley et al., 1983, 1987a; Polak et al., 1989). The pattern seems to be loosely based on the molecular size and shape of the odorant. T h e same kind of approach has been tried iti z~tvousing behavioral measures of response (Mason et al., 1985, 1987a,b). Although there are not enough data to show a pattern, some selectivity is apparent. 2 . Candzdate Receptor Molecules An “anisole-binding” protein has been purified from a detergent extract of dog olfactory epithelium (Goldberg et al., 1979; Price and Turpin, 1980). Price and Willey (1 987) report a “benzaldehyde-binding” protein similarly purified. Both proteins are present in olfactory tissue but not in respiratory tissue. They are distinguishable from each other but both are 61 kDa. Antibodies to either can suppress the electrophysiological response of the olfactory tissue. Price and Willey (1988), using a number o f monoclonal antibodies to these proteins, have demonstrated some odor-selective suppressions of response that seem to parallel the binding properties of the proteins. Fesenko et al. (1987, 1988)have also advanced candidates for olfactory receptor molecules. ‘These can be extracted from rat olfactory tissue with nonionic detergent. Odorant-binding activity is found in a heterogeneous glycoprotein of 88 kDa that is associated with a glycoprotein of 55 kDa. Purified antibodies to the 140-kDa complex bound only to olfactory tissue and were capable of abolishing electrophysiological responses. A 98/.56-kDa protein complex has also been isolated from the olfactory epithelium of the skate (Novoselov et al., l988a,b). This will bind amino acids that are olfactory stimuli for this fish and shows enhanced GTPase activity in their presence and could possibly be a recept o r 4 protein complex. Lancet’s group have described a heterogeneous 95-kDa glycoprotein extractable from preparations of frog olfactory cilia (Chen and Lancet, 1984; Chen et a/., 198613). Bovine olfactory cilia also contain specific transmembrane glycoproteins and phosphoproteins (Kropf et al., 1987). There is evidence for the existence of proteinaceous odorant-binding sites in the epithelium. Microwave radiation causes a decrease in the epithelium’s ability to bind camphor but has no effect on the binding ability of a detergent extract, suggesting that proteins responsible for binding are being shed (Phillippova et al., 1988). T h e binding of an

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odorous steroid can be directly measured (Persaud et al., 1988b) and the odorant-binding protein (Section VI,E) was originally detected by its ability to bind odorants.

B. RECEPTORCELLS

1. Response to Odor Concentration Any individual cell shows a sharp increase of spike rate with concentration over a fairly narrow concentration range (Getchell and Shepherd, 1978; van Drongelen, 1978; Trotier and MacLeod, 1983; Getchell, 1986; Lidow et al., 1987b; Firestein and Werblin, 1989). However, the population as a whole shows a more gradual increase as more cells are recruited to the response, until, at very high concentrations, almost all cells respond (Duchamp-Viret et al., 1989). It has been suggested (Lidow et al., 1987b) that an individual cell may only be transmitting a useful message over a narrow concentration range, with different cells covering different parts of the total range. Using intracellular recording techniques, Trotier and MacLeod (1983) showed that a receptor cell ceases to generate spikes during repolarization even if the intracellular potential is positive with respect to the threshold. This implies that a cell signals some degree of information about rate of change of concentration.

2 . Tuning In aquatic invertebrates there is evidence that some receptor cells display fairly sharp tuning (Fuzessery et al., 1978;Johnson and Atema, 1983; Derby et al., 1984). The lobster has cells sensitive to the adenine nucleotides; the electrophysiological response spectrum of some parallels that of the P, purinoceptor (Derby et al., 1987),and that of others is like that of the P, purinoceptor (Carr et al., 1986, 1987).The lobster also has cells responsive to hydroxyproline. Again there are two types of cells (Johnson et al., 1987, 1988a), one being most responsive to hydroxyproline and the other being most responsive to gelatin, a hydoxyproline-rich polypeptide. This pattern of tight tuning of receptor cells in the water breathers may have much to do with the nature of their chemical environment (e.g., Carr, 1987). The pattern may be lost when complex stimuli are used (Derby and Ache, 1984b; Giradot and Derby, 1988) and it has not been found among the air-breathing animals. Studies by Revial et al. (1982, 1983), Sicard and Holley (1984), and Sicard (1985) on frogs show that the cells are very responsive. At high

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odor concentrations, about 50% of cell-odor combinations show a response. Although there is some pattern of cellular selectivity toward different odorants, seemingly based on odorant shape and size, there is no clear grouping of cells into distinct types. During development in the rat, cells first become responsive to odor generally and gain selectivity for odorants later (Gesteland et al., 1982). This selectivity seems to be acquired in parallel with the appearance of olfactory cilia (Menco, 1985; Menco and Farbman, 1985) and an increase in the number of particles in the sensory membranes (Menco, 1988a). In the frog, cells d o not seem to go through a phase of general sensitivity (Lidow et al., 1987a,b). Certain antibodies can recognize subclasses of olfactory receptor cells (Allen and Akerson, 1985a,b; Fujitd et al., 1985; Mollicone et al., 1985; Mori ct d.,1985; Mori, 1987b; Schwob and Gottlieb, 1987, 1988; Akerson, 1988), but there is no evidence that any of these subclasses correspond with functional classes. There is indication that at least some of them may represent developmental states (Onada, 1988a,b; Astic et al., 1989; V. M. Carr et al., 1989). It may also be possible to classify cells by their carbonic anhydrase content (Brown rt al., 1984),but there has been no demonstration of a correlation with odor reception properties. 3 . Patterns of Responses

T h e epithelium does not show a uniform responsivity toward odorants and displays a different pattern of sensitivity toward different odorants (Moulton, 1976; Kubie et al., 1980; Mackay-Sim and Kubie, 1981; Mackay-Sim et al., 1982; Mackay-Sim and Shaman, 1984; Edwards et al., 1988; Triotskaya, 1988). T h e response pattern for a particular odorant is stable as the odor concentration is raised (Mackay-Sim and Shaman, 1984), although “best” areas may reach a maximum response while “poor” ones catch up as the concentration increases. Although the differences in patterns for different odorants are not large, they do indicate some degree of clustering of alike receptors.

C. RESPONSES IN THE OLFACTORY BULB 1. El~ctrophysiolopcal Studies

Patterns of activity in the olfactory bulb d o not relate solely to the stimulating odor. Information about the phase of the respiratory cycle is present in rabbits (Chaput and Holley, 1985; Chaput, 1986; Chaput and Lankeet, 1987) and in neonatal rats (Mair and Gesteland, 1982). ‘This

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seems to arise by an odorant’s ability to increase discharge rates on inspiration and decrease them on expiration. However, there seems to be no “gating” of information in the higher centers, as animals are able to detect and discriminate different patterns of electrical stimulation of the bulb regardless of the phase of the respiratory cycle (Monod et al., 1989). Responses in the bulb also depend on the animal’s previous experience with odor (Section IV,C,4) and possibly on air movement (Potapov, 1987). It is suggested (Hamilton and Kauer, 1985, 1987, 1988, 1989; Kauer and Hamilton, 1987) that the temporal pattern of activity elicited in a mitral or tufted cell by an odor pulse depends on the bulbar circuits and does not simply reflect the underlying activity of the receptor cells. The initial response of a mitral or tufted cell to an odor pulse is hyperpolarization; this may (depending on the particular odorant) be followed by depolarization and there is also a late hyperpolarization. According to the hypothesis, odor stimulates widely scattered epithelial receptor cells (Kauer and Moulton, 1974), which excite periglomerular cells that in turn inhibit mitral or tufted cells, giving rise to the initial hyperpolarization. More tightly clustered cells in the epithelium are connected directly to mitral or tufted cells. Depending on the type@)of receptor cell involved, an odor pulse may result in the depolarization of the mitral or tufted cell but the information is transmitted more slowly and, for moderate odor concentrations, the depolarization follows the initial hyperpolarization. The late hyperpolarization is mediated by the granule cells. a. Responses with Odor Concentration. An odorant may cause excitation or suppression of the activity of a bulbar neuron. There is disagreement about whether the kind of response evoked and the temporal pattern of firing are stable as the concentration of the odorant is varied. Harrison and Scott (1986) found that cells can have a complex variation of response type with odor concentration, that two cells may show similar or different variations of response with concentration, and that it is not possible to classify such response profiles. Wellis et al. (1989) emphasized that cells display complex patterns of firing and that these patterns change with stimulus type and intensity. Kauer (1974) found both excitatory and suppressive responses and hypothesized that excitation might change to suppression as odor concentration rose, and Duchamp (1982) found that suppressive responses from bulbar cells were most often seen at high stimulus concentrations. Kauer ( 1974) hypothesized that excitations coded for stimulus intensity and that different cells might be excited by different parts of the concentration range. This concept of concentration tuning has found support in the work of Mathews (1972), Shibuya et al. (1977),Daval and

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Leveteau (1982), Meredith (1986), Reinken and Schmidt (1986), and Hamilton and Kauer (1989). T h e general finding is that for any given odor and cell, response type rarely changes with concentration, whereas response magnitude may vary with concentration, there often being an optimum concentration range. There can be changes in the temporal firing pattern of a neuron in response to odorant stimulation (e.g., Mair, 1982a; Meredith and Moulton, 1978). Meredith and Moulton (1978) attribute many of the changes in the temporal firing pattern of cells to the fact that the stimulus concentration in the mucus changes with time. Duchamp and Sicard (1984) found that the secondary cells are less sensitive to concentration than are the primaries, and Duchamp-Viret et al. (1989),emphasiLing the importance of maintaining the integrity of all the epithelial connections to the bulb, found that if a bulbar neuron responded at a certain concentration then it would respond at all higher concentrations. At high concentrations most of the bulbar cells would respond. Dgving (1987), using stimuli whose profiles were ramps rather than pulses, found both excitatory and inhibitory responses with little change of response character with concentration and noted that the majority of cells were unresponsive to most odorants even at high concentrations. Mair (1982b) has studied the adaptive properties of rat bulbar neurons. Some neurons change excitability if repeatedly challenged with odor, but the kind of change (increase or decrease) is characteristic of the neuron and not of the odorants used nor their concentration. Preexposure may change the extent of response but not the temporal firing pattern. Schild and Zippel (1986) have studied bulbar responses to repeated odor stimulation in fish. Usually, responses were stable but in a few cases there was a more pronounced response to later presentations. b. Tuning. There are differences of opinion as to the extent to which a buibar cell can be said to be tuned to particular odorants. Harrison and Scott (1986) found that the response type changed in a complex fashion with both the nature of the stimulus and its concentration. However, Duchamp (1982) and Duchamp and Sicard (1984) found that bulb cells were better able to discriminate between odorants than were receptor cells and thus were more tightly tuned. Hamilton and Kauer (1989) have shown that different cells are differentially activated by different odorants. T h e hypothesis is that it is this pattern of activation that encodes odor quality and that the patterns arise as different cells have different inputs from the epithelium (see also Kauer, 1987). Most other authors have not used a wide enough range of odorants to investigate the tuning of the cells. Mair (1982a) noted that odorants differed in their

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ability to excite a particular cell and that cells differed in their ability to be excited by a particular odorant. 2. Optical Studies Events in the olfactory bulb have been studied using voltage-sensitive dyes and optical monitoring (Orbach and Cohen, 1983). The technique can follow events occurring on a tens of milliseconds time scale and may be very useful in a system wherein patterns of differential activation may be more important than the activation of any one cell. It has been applied using electrical stimulation (Kauer, 1988) to show the spread of activity through the bulb and long-lasting activity in the granule and mitral cells. It has also been used with odor stimulation (Kauer et al., 1987) to show that there is a great heterogeneity of response across the bulb, indicating the activation of parallel circuits. The general pattern is of a rapid rise in activity following the start of an odor pulse, followed by a slower return to baseline while the odor is still present in the olfactory mucus. 3. 2-Deoxyglucose Studies Early studies of the uptake of 2-deoxyglucose (2DG) by the olfactory bulb during odor stimulation showed that each glomerulus tends to label uniformly but that there are differences between glomeruli. Each glomerulus may therefore be a functional unit (Sharp et al., 1977; Skeen, 1977; Stewart et al., 1979; Lancet et al., 1982). Most of the metabolic activity responsible for label uptake is presynaptic (Benson et al., 1985). Different odorants induce different patterns of labeling Uourdan et al., 1980; Penderson et al., 1986a). Royet et al. (1987), using computer-assisted image analysis, have shown that exposure of mice to isovaleric acid and to amyl acetate vapors cause 2DG labeling in different patterns of glomeruli. A strain of mouse reported to have impaired ability to detect isovaleric acid shows some changes in the acid-induced pattern relative to the controls but a normal pattern in response to the ester (Sicard et al., 1989). Hypogonadal mice are reported to have an impaired sense of smell, and a 2DG study by McQueen et al. (1988) showed that odor could stimulate metabolism in the sensory cells of these animals but not in the glomeruli. Propionic acid vapor induces 2DG labeling in a fairly well-defined cluster of glomeruli. This part of the olfactory bulb can be removed surgically with little or no impairment of the animal’s ability to detect or respond to propionic acid or other stimuli (Slotnick et al., 1987). This may simply reflect the redundancy present in the olfactory system (e.g.,

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S. G. SHIRLEI:

Risser and Slotnick, 1985). But Coopersmith and Leon (1989) have pointed out that in this tissue there might be a significant contribution to 2DG uptake from the generation of the reducing power needed to drive a detoxification system. 2DG labeling experiments are performed with prolonged exposure to high odorant concentrations and it is possible that some labeling represents detoxification rather than neural activity. It is not clear why a protective mechanism should lead to differential labeling of glomeruli, but the finding of Slotnick et al. (1989) that labeling patterns persist at very much reduced odor exposures does give confidence that the 2DG method produces results relating to stimulation rather than adaptation or protection. Astic et al. (1988) have found that as far as 2DG uptake is concerned, cells do not adapt rapidly and totally. 4. Leu rning

It is clear that responses in the olfactory bulb do not relate solely to the nature and intensity of the stimulus but are largely conditioned by learning. In neonatal rats, experience with an odor coupled with tactile stimulation causes changes in the bulb such that there is enhanced uptake of 2DC; specifically into those glomeruli that normally label in response to that odorant (Coopersmith and Leon, 1984; Coopersmith et nl., 1986; Sullivan and Leon, 1986). The effect of this early training persists into adult life (Coopersmith and Leon, 1986).Such training can also cause changes in single-unit responses toward the odorant (Wilson and Leon, 1986; Reinken and Schmidt, 1987; Wilson et al., 1987). These results have been summarized by Leon (1987). T h e learning of a significant odor is accompanied by a change in the bulbar EEG that the odor elicits (Freeman and Schneider, 1982; Freeman and Grajski, 1987; Freeman and Baird, 1987). There is a rich projection of noradrenergic neurons from the locus coeruleus to the granule cells of the olfactory bulb (Shipley et al., 1985), and noradrenalin has been shown to be important in the learning of olfactory cues in adult mice (Rosser and Keverne, 1985; Rosser et al., 1986; Dickinson and Keverne, 1988), rabbits (Gray et al., 1986), and sheep (Pissonier et al., 1986) and also in infant rats (Sullivan et al., 1989). Gervais (1987) has demonstrated that y-aminobutyric acid (GABA), released by numerous bulbar cells on odor stimulation, enhances the release of noradrenalin in olfactory bulb slices. T h e role of noradrenalin in olfactory learning has been reviewed by Gervais et al. (1988). It is possible that the noradrenergic projections are signaling the animal’s state of vigilance (Aston-Jones, 1985), thus setting the time at which a memory should be formed, a view comparable with the findings of Kaba and

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Keverne (1988) and Kaba et al. (1989) on the effect of GABA and noradrenalin blockers.

V. Transfer of Information

It may now be possible to fit the various parts of the olfactory system into perspective and follow the flow of information through the system. This section is written within the framework of the vectorial concept of odor recognition and of an adenylate cyclase-coupled mechanism, but some of the remarks remain valid outside those frameworks. A. THEVECTORIAL REPRESENTATION The “vectorial” model of odorant recognition has been given a formalism by Schild (1988) and the concept can be seen in previous works (Amoore, 1970; Polak, 1973; Shirley, 1984; Lancet, 1986) and, implicitly o r explicitly, in the works of many other authors. The basic principle is simple. An odorant will stimulate not one but several types of receptors (molecule or cell). The degree to which it stimulates a particular type of receptor molecule depends on the nature of the receptor and nature and concentration of the odorant. Recognition depends on the evaluation of the relative degrees of stimulation. At its simplest, the vectorial representation assumes that the interaction between an odorant and any one type of receptor molecule is characterized by a dissociation constant (or other characteristic parameter with the dimensions of concentration) that is determined by the geometry of the binding site and the molecular properties of the odorant. This constant may, of course, be modified by the presence of a linked transductory system, e.g., by guanine nucleotides. The dissociation constants displayed by any one type of receptor toward different odorants should constitute a “classical” structure-activity relationship intelligible in molecular terms. That the (airbreathing) olfactory system tends to display rather diffuse and ill-defined structure-activity relationships simply reflects the rather broad tuning of the receptors and that odor recognition involves events occurring at more than one type of site. If the interactions between an odorant and the various receptors can each be described by a single dissociation constant, then as far as the

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system is concerned any concentration of the odorant can be represented by a vector whose components are the ratios of concentration to the various dissociation constants. T h e length of this vector relates to the concentration, and the direction of the vector is characteristic of the odorant. If different odorants simply compete for receptors and receptors simply report the degree of occupancy, then any mixture of odorants can be represented by a vector that is the sum of the individual vectors. T h e vectorial model does not assume that the coding of the vector or its components is linear o r simple, o r that all the information in the vector is of biological relevance, rather only that the vector describing the odorous content of the mucus and its variation with time is the only information available to the system. In essence, the olfactory system exists in order to extract the biologically relevant information from this vector. In the vectorial model, odor recognition is a system property-information from many sources (types of receptor molecules) must be collated. In some ways this is analogous to the color vision system; the perceived sensation depends on the relative degree of stimulation of several receptors, and activation of the “green” receptor does not of necessity produce the sensation green. This contrasts sharply with the situation in the pheromonal system of insects, in at least part of the olfactory systems of fish and, possibly, in the vomeronasal system of mammals, wherein recognition, although dependent on some further data processing, is largely a property of the receptor molecule. The vectorial model does not deny the possibility that a particular odorant may stimulate only one type of receptor molecule and so be “primary” in the sense of Amoore (1970), o r that one particular type of receptor molecule may respond to only one compound and so be able to recognize an odorant unaided by the rest of the system. Rather it contains these possibilities as (probably rare) special cases.

B. ODORCONCENTRATION AND SIGNAL/NOISE The absolute concentration of an odorant in the environment is, in general, a parameter that carries rather little information to an animal. Atmospheric conditions will tend to change absolute concentrations unpredictably over a wide range. Much more information is carried by concentration changes and gradients and by odor quality, which will tend to be little perturbed by atmospheric conditions. Overall, the olfactory system is poorly sensitive to stimulus concentration. This may sim-

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ply reflect the natural lack of information carried by this part of the incoming signal and it does not imply that concentration is unimportant at the periphery of the system. In the vectorial model, within a single sniff, concentration, and in particular concentration increase, are very important, as they dictate the degree of stimulation that an odorant will initially produce in all the different types of receptor molecules, and this concentration information must be preserved at least to the point in the system where the “information streams” from each type of receptor recombine. Any real system must cope with the problem of noise. The olfactory mucosa is not intrinsically odor free-it is exposed to odor from the animal, to volatiles circulating in the blood, and to persistent environmental odorants; even permanent gases such as nitrogen are odorous if their concentration is changed (Laffort and Gortan, 1987). Although there is what might be described as a noise reduction system (Section VI,D), it is unlikely to be perfect. The olfactory system must be able to discriminate a real (environmental) signal from this chemical noise. When an odorant partitions between air and an aqueous medium, it is concentrated into the liquid (Section V1,A). A receptor cell may be triggered by the opening of a single ion gate (Section III,A,l). This means that, in principle, a single cell carrying a single ion channel gated by an odorant in the nanomolar concentration range is sufficient to explain the sensitivity that the olfactory system displays toward many compounds. T h e olfactory system is many orders of magnitude more extensive than this. This may not be a strategy for increasing sensitivity per se, although it will do so. In the face of a constant level of odorant, the receptors will themselves produce noise because, at any given instant, a particular receptor molecule must be bound or not and a particular cell must be spiking o r not. The use of a large number of receptors allows this noise to be averaged out so that the system can faithfully detect significant changes in a possibly high background. The background itself may also be quite variable because it is largely of biological origin, and thus unlike Brownian motion, which ultimately sets an irreducible noise level for hearing. Such variations in background could partially explain the notorious variability of olfactory detection threshold measurements and the finding of Stevens et al. (1988) that a large part of this variability is between measurements made on the same subject at different times rather than between measurements made on different subjects. The threshold measurement could be reporting the system’s ability to discriminate a signal from the prevailing background rather than some absolute molecular or cellular property and thus yields an answer that changed with the background. It has been noted (Handrich and Atema,

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1987; Voigt and Atema, 1987) that chemical signal-to-noise ratios are important in electrophysiological and behavioral responses in taste, and this may also be true in olfaction.

c. INFORMATION TRANSFER FROM M U C U S TO RECEPTORMOLECULE I . Imperfectzons As the vectorial model conceives odor quality to reside in the ability of a stimulus to activate different receptors differentially, it is important that such differences are not capriciously altered. If one species of receptor molecule were to saturate, then its output would remain invariant while those of others were free to change with concentration. This would cause a concentration-dependent change in perceived odor quality. It is likely that the affinities of receptor-odor interactions are generally fairly weak, with dissociation constants typically in the micro- to millimolar concentration range rather than the nanomolar range common for neurotransmitter and hormone receptors. This conclusion is reinforced by the general difficulty in demonstrating saturation of olfactory responses. The olfactory system does exhibit concentration-dependent changes of perceived quality (e.g., Gross-Isseroff and Lancet, 1988),although these rarely seem t o be of the degree necessary to cause niisrecognition of an odorant. Such changes are easier to explain in a vectorial system than in one that relies on dedicated receptors to recognize odorants, where, if the receptor saturates, only quantitative information is lost. Other phenomena that could lead to concentration-dependent quality changes are allosteric binding, synergy o r antagonism in the binding of mixture components, and partial agonism. If synergy and antagonism were common and extensive then receptor occupancy in response to a mixture of arbitrarily chosen components would become chaotic-unpredictable from a knowledge of the individual bindings. 2. Mz.xture.y

The olfactory system has to deal with mixtures. In the vectorial model, it is important to distinguish the case wherein one odorant is presented in the preexisting background of another, so the new arrival can be recognised by, essentially, a vector subtraction, from the case wherein the components of a mixture are presented simultaneously. These may result in the presence of exactly the same odorant molecules in the olfactory mucus but a completely different sensation. It is also important to bear in mind that there are physicochemical processes (Sec-

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tion V1,A) such that the simultaneous presentation of the components of a mixture at the naris does not necessarily mean that all components will arrive simultaneously at all points of the epithelium and that the timing of such delays could carry meaningful information. In the vectorial model, different components of a mixture can compete for a particular receptor. Interactions other than competition (synergy o r suppression) are undesirable as they will lead to concentrationdependent changes of odor quality. In a model wherein each component of a mixture interacts only with its own receptor, there should be independence of response at the periphery of the system. Mixtures have been used as stimuli for invertebrates and many authors have found evidence for complex peripheral interactions between mixture components (Derby and Ache, 1984a; Zimmer-Faust et al., 1984; Derby et al., 1985; Gleeson and Ache, 1985;Johnson et al., 1985; Borroni et al., 1986; Carr and Derby, 1986a,b; McClintock and Ache, 1989b). However, in the catfish, Caprio et al. (1989) find that peripheral electrophysiological responses to mixtures are predictable from the responses to the individual components simply on the assumption of competition. There has been very little work on peripheral responses to mixtures in mammals. Bell et al. (1987) report that the bulbar 2DG labeling pattern characteristic of an odorant disappears when the compound is presented as an undetectable component of a mixture. Mixture responses and their predictability are good indicators of the nature of the olfactory system: dedicated receptor, vectorial, imperfect vectorial, or chaotic. It is to be hoped that such work will be extended to the air-breathing vertebrates. 3. Receptor and Receptorless Systems There is some indication that the mammalian olfactory system has proteinaceous receptor molecules (Shirley et al., 1983), and there are some candidate receptor molecules (Section IV,A,P), and strong indications that some invertebrates and fish have receptor molecules. But until there is a rigorous demonstration that a molecule is, in fact, a receptor, it must be remembered that discrete receptors are not necessary. Lipids membranes can respond to odorants (Section II,D), and lipids of different composition can respond differentially to different odorants; this is a large part of the requirement for a system working on the vectorial principle. It is difficult to see how the lipid composition of a membrane could be controlled with sufficient precision to give the long-term stability of response that might be expected of a protein molecule, and, until proteinaceous receptors are positively identified, this is the main argument against receptorless systems (Shirley and Robinson, 1988). Assuming the existence of proteinaceous receptors, the problem is to

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explain a requirement of the vectorial model: that any odorant capable of binding to a receptor will cause the same kind of response. Receptors of the neurotransmitter or hormonal pattern often have a larger number of antagonists than agonists, as the conditions for a molecule to bind to a receptor are less stringent than those for it to bind and stimulate. One of the characteristics of the olfactory system is the striking absence of antagonists. There are clear evolutionary advantages of a sensory system whose receptors remain functional whatever the incoming stimulus, but this behavior places restrictions on the chemical mechanisms of odor-receptor interaction. A possible mechanism in the olfactory case is that both binding and stimulation are primarily hydrophobic and both depend on an odorant’s ability to exclude water from the binding site, but the odor-receptor interaction requires extensive study. 4. Broadlj and Narrowly Tuned Receptors

The vectorial representation makes no assumption about the breadth of tuning of the receptor molecules; the only assumption is that overlap between the various response spectra is the rule rather than the exception. There is chemical modification evidence that at least some receptors are broadly tuned (Section IV,A,l) and an argument based on a reciprocity relationship and the fact that second-messenger changes are observable. Draw u p a table containing all n possible odorants as columns and all rn types of receptor as rows. Mark those points of intersection that correspond to interactions sufficiently strong to be of practical significance; there will be q marks. A randomly chosen receptor will, therefore, be stimulated by, on average, qlm odorants or a fraction ql(m x n) of all odorants. A randomly chosen odorant will on average stimulate qln receptors or a fraction ql(m x n) of all receptors. If the “average” receptor is stimulated by x% of all odorants, then the “average” odorant stimulates x’jT of all receptor types. ‘The second-messenger enzymes must be compartmentalized between cells and possibly between different compartments in the same cell. ’10 allow two different types of receptor to stimulate the same part of the second-messenger pool would add two different components of the vector, destroying the information needed for odor recognition. N o subsequent part of the system could determine which receptor had been stimulated. (A rotation of the coordinate system of the vector, which would preserve the information, requires both positive and negative coefficients, i.e., one receptor to stimulate and the other to inhibit. To date, odorants have only been observed to stimulate adenylate cyclase.) If receptors were tightly tuned, say a typical receptor responded to only one type of odorant in 1000, then a typical odorant would stimulate only

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one type of receptor in 1000 and activate only 1/1000 of the adenylate cyclase. Such an activation would not be detectable above the total basal activity, as ligand-activated/basal activity ratios for any part of the pool are unlikely to exceed a factor of about 20. But, typically, single-odorant activations of the cyclase are 30-150% of the combined basal activity. That cyclase activations are detectable at all and certainly that they are of this magnitude are strong arguments for broad tuning. Broadly tuned receptor molecules with overlapping response spectra guarantee that virtually any compound will be detectable. Faced with a novel stimulus, an animal may learn its significance but has no need to “learn” to detect it by inducing new types of molecules. 5. Receptor Heterogeneity In general, different types of receptors will display different affinities toward the same odorant. This is the sense in which heterogeneity is used here; the receptors may, of course, be heterogeneous in other senses, such as glycosylation, as well. As currently measured, the response of adenylate cyclase to odorant is the sum of activations due to the individual receptor types. This sum will vary with the concentration of the odorant and the heterogeneity will cause the sum to display a smaller Hill coefficient than any one component actually has. The observation that adenylate cyclase activation has a Hill coefficient < 1 does not imply that the interaction of an odorant with any individual type of receptor is nonhyperbolic. Indeed, concentration-activation curves can easily be fitted as the sum of hyperbolae (S. G. Shirley, unpublished observations). Receptor heterogeneity may be the underlying reason for the difference between the Hill coefficients of individual and summated measures of peripheral responses. Summated measures, e.g., adenylate cyclase activation (Pace et al., 1985; Shirley et al., 1986), electroolfactogram amplitude (a generator current measure) (Ottoson, 1956; Silver, 1982; Shirley et al., 1987b), and spike recordings from nerve bundles (Silver, 1982), show low Hill coefficients or power law exponents. Individual responses, e.g., single-cell receptor potentials (Trotier and MacLeod, 1983) or single-cell spike rates (Getchell and Shepherd, 1978; Trotier and MacLeod, 1983; Lidow et al., 1987b; Firestein and Werblin, 1989), show a much sharper concentration dependence.

6 . Classijication With broadly tuned receptors whose response spectra overlap, it is not satisfactory to attempt to classify a receptor simply on the basis of whether or not it responds to a particular odorant. Some measure of the interaction strength, such as dissociation constant, must be used because

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it is likely that, at high enough concentration, a very large number of odorants may be capable of stimulation.

D. INFORMATIONTRANSFER FROM RECEPTOR MOLECULETO CELL 1. S’etidzziitj

T h e sensitivity of a receptor cell does not depend solely on the dissociation constants of its receptor molecule(s). T h e number of individual receptor molecules will affect sensitivity. An array of several hundred thousand receptor molecules (the number of particles per cell seen in freeze-etch electron microscopy), each with a dissociation constant of 1W4M , is more sensitive than a single molecule of dissociation constant 10-9 M , has a much wider dynamic range, and is potentially less noisy because events at many sites can be averaged. T h e question of the number of copies and stoichiometry of the various kinds of molecules is important and intimately related to the question of sensitivity. Currently it is not known whether the transductory apparatus is a “funnel” transferring information from a large receptor array to a few cyclase molecules and ion gates and so continuing the pattern of convergence seen at higher levels of the system, or an “amplifier” opening many ion gates for a single event at a receptor. 2. Adaptatzon It is also possible that the transductory apparatus is not static. Though it is unlikely that a receptor cell could internalize significant numbers of receptor molecules on the time scale of tens of hundreds of milliseconds typical of transduction, the possibility that this mechanism might function as a long-term sensitivity control should not be overlooked. T h e degree of coupling between receptor and cyclase is not necessarily fixed. T h e finding that most of the cyclase in cilia1 preparations is not coupled to the G protein indicates this coupling is potentially a mechanism for changing the cell’s sensitivity. The action of the calcium ion on the second-messenger enzymes can change the sensitivity of the cell. T h e opening of ion gates to change the resting potential and impedance of the cell will also change sensitivity and there is the possibility (Section VI,C) of some measure of central control of the cells. Adaptation o r the ability of the cell to adjust its sensitivity according to prevailing conditions is a prominent feature of the olfactory system and it will require much effort to elucidate the relative importance and the time scales of the various possible mechanisms. Sensitivity and adap-

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tation are intimately bound up with edge enhancement, or the cell’s ability to respond strongly to increases in stimulus concentration. Again there are several possible parts of the mechanism that could contribute to this: the action of calcium on second-messenger enzymes and ion channels, the possibility that odorants may control two kinds of ion channel (assuming correct timing) (Persaud et al., 1988a), and the possibility that rate of change of depolarization has an influence on spike generation (Trotier and MacLeod, 1983). That a cell should be particularly responsive to odor concentration changes is not surprising, because dogs (Neuhaus, 1981), rabbits (Karpov, 1980), and humans (Laing, 1986) can make decisions about odors within the time taken for a single sniff. Within this time (150-450 msec, depending on species), the concentration of odorant within the mucus will be changing (Section V1,A). A cell that is highly adapting and emphasizing concentration increases could, in vivo, be receiving absolute concentration information at its receptor molecules but processing it to output the far more useful information about concentration changes. Under experimental conditions with a relatively constant background and the use of odor-pulse stimulation, this will still be true, but under those conditions, this output has a fairly simple relationship to the pulse concentration. The view of receptor cells as concentration sensors or detectors may simply reflect those particular experimental conditions. Much work needs to be done to understand the extent to which the adaptive processes extract biologically relevant information from the cell’s input. In highly adaptive cells, the concept of dynamic range needs careful consideration. The cell may remain responsive over a wide range of background concentration but may only respond over a small range of concentration increase. Activation of currents by cyclic AMP or analogs also has a Hill coefficient > 1 (Nakamura and Gold, 1987; Persaud et al., 1988a; DeSimone et al., 1988) and the relationship between depolarizing current and spike rate is sharp (Hedlund et al., 1987; Masukawa et al., 1987). This suggests that any particular cell signals useful information over a quite narrow concentration range, as suggested by Lidow et al. (1987b), although it may remain maximally activated at higher concentrations (Duchamp-Viret et al., 1989). But there is no reason to suppose that this range stays constant in the face of a changing background. 3. The Arrangement of Receptor Molecules on Cells

A key question is of the arrangement of receptor molecules on cells, because this arrangement relates the response spectra of the molecules to the response spectra of the cells. The response spectra of cells are stable and tend to be broad, but have so far resisted attempts at classifica-

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tion. 'The remarks of Section V,C,S apply to the classification of cells as well as to the classification of molecules, with the added complication that cells may show variations of sensitivity that are not of receptor molecule origin. Perhaps the classification scheme most likely to succeed in these circumstances is one based on the ratios of the concentrations at which two or more different odorants elicit the same response. This kind of approach has been used successfully to classify olfactory cells in insects (Kafka, 1987). Ion gates, adenylate cyclase, G protein, and, presumably, receptors are membrane-bound molecules and as there are diffusion barriers in the membrane these components could be compartmentalized within the cell. Cyclic AMP is soluble, but on the time scale of transduction could only difFuse for a distance of a few micrometers, not, for instance, for the entire length of a cilium. So it is likely that any given ion gate responds only to events occurring at local receptor molecules and conceivable that different kinds of receptor molecule could be compartmentalized within the same cell to control different ion gates. But neither odorants nor second messengers could diffuse from the trdnsductory apparatus to the cell body at the speed of transduction, and this signaling is believed to be purely electrical. T h e spike-generating apparatus receives only a summated signal from the receptor molecules. If a cell carries more than one type of receptor molecule and odor-quality information resides in the differences in an odorant's ability to stimulate the different receptor types, then addition of the signals will destroy the infbrniation. If a cell carries more than one kind of receptor molecule, then the mechanism must involve both addition and subtraction and it will be necessary to demonstrate odor-specific excitations and inhibitions occurring within the same cell. Within the olfactory bulb, there is ample evidence for both excitatory and inhibitory pathways and no conceptual difficulty in understanding the extraction of qualitative information.

4. Conuergence T h e effect of the high degree of convergence between receptor and secondary cells has been discussed by van Drongelen et al. (1978), who pointed out that the secondary cells should be more sensitive to odorants than are the receptors. If a function of the secondary cells is to average noisy signals, then the sensitivity should improve roughly with the square root of the number of receptors connected to each secondary. For a convergence ratio of about 1000, the sensitivity of a secondary cell should be about one and a half orders of magnitude greater than a receptor. This is close to what has been observed by Duchamp-Viret et al.

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(1989). These authors also point out that a bulb can respond to odorants with many of its peripheral connections broken, but that its sensitivity is drastically reduced under these conditions.

VI. Perireceptor Events

T h e events that occur in the olfactory mucus prior to, during, and after transduction are of great significance and have been reviewed by Getchell et al. (1984).

A. THEOLFACTORY Mucus Very little is known about the olfactory mucus. The mucus over the respiratory region has two layers of different consistency, but that over the olfactory region is a single layer (Menco, 1989b). A histochemical study by Gladysheva et al. (1986) revealed acidic, sulfated, and neutral mucopolysaccharides in vertebrate mucus. T h e mucus is also sodium poor and potassium rich compared with most intercellular fluid (Joshi et al., 1987). Hornung et al. (1987) have studied the equilibrium partitioning of odorants between air and the olfactory mucosa. It must be emphasized that the mucus (which bathes the receptor apparatus) is not the major component of the mucosa and that dynamic considerations (deviations from equilibrium) may be important in viva However, the conclusion of these authors is that for most odorants the mucosa/air partition coefficient is within a factor of two or three of the water/air partition coefficient, which can therefore be used as a rough guide. As the concentration of an odorant partitioning into the mucus is of fundamental importance to an olfactory biochemist studying receptor affinities, etc., it is worth reiterating the physical chemistry of the situation. T h e equilibrium distribution of an odorant between water and air is usually described by a partition coefficient (molar concentration in water/molar concentration in air) or a Henry coefficient (the limit at low concentration of the mole fraction in air/mole fraction in water). Both are temperature sensitive, as is the ratio of molar volumes of water and air that relates the two. Typically, odorants have partition coefficients of between 50 and 10,000, although examples can be found outside this range, but for virtually all odorants the molar concentration in water is higher than that in air. By elementary thermodynamics, the air above a

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saturated solution of a sparingly soluble material contains vapor at the saturated vapor pressure. So a knowledge of the saturated vapor pressure, which is related to the molar concentration in air, and the solubility allows an estimate of the partition coefficient if no better data are available. 'The presence of a cosolute in the water affects the distribution only insofar as it changes the solution properties (dielectric constant or structure, etc.) of the water. i n this respect, probably the most significant solutes in mucus are the salts, and saltwater/air partition coefficients should provide a better approximation to the olfactory situation. For biochemical purposes, the significant variable is the concentration of material free in solution, because this is the thermodynamic driving force determining binding to a receptor and it is this concentration that is related via the partition coefficient to the concentration of vapor above the liquid. The ability of a cosolute to bind odorant is irrelevant, as it simply creates a third compartment whose contents come to equilibrium without disturbing the relationship between free solution concentration and vapor concentration. T h e same is true for any other entity that can absorb odorant. At equilibrium, the free solution concentration is simply related to the vapor concentration despite binding, the solution of odorant into membranes, and monolayers formed on equipment surfaces, etc. Such compartments do affect the time taken to reach equilibrium when the tissue is stimulated via the gas phase, as more material must be transferred. But the finding that mucosa/air partition coefficients are fairly close to water/air coefficients means that this delay is neither large nor unpredictable. The olfactory receptor molecules are very superficial in the mucosa and the mucus at their depth will equilibrate with the air more quickly than will deeper material. if the tissue is stimulated by an odorant in moving air, then the time to achieve equilibrium should be little greater than the longer of two characteristic times. T h e first is the time for an odorant to diffuse to equilibrium in the mucus (tens to a few hundred milliseconds, depending on the mucus thickness). The second is the time for sufficient air to pass over the tissue to deliver enough material to bring the superficial layers to their equilibrium concentration; this time depends on the partition coefficient of the odorant, as this determines how much material must be transferred, but under typical experimental conditions and for odorants of partition coefficients up to a few thousand, this time should not exceed 1 sec. So for odorants of low and moderate partition coefficients, the free solution concentration in the vicinity of the receptor molecules should, under experimental conditions on a time scale of seconds, be close to the equilibrium value. But during a sniff zn vivo, on a 100-msec time scale, concentrations are likely

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to be changing. Prediction of these changes requires detailed knowledge of the aerodynamics of the cavity and physical chemistry of the particular odorant. Odorants are delivered to the olfactory epithelium in a moving airstream that passes over the relatively stationary mucus layer. This, coupled with the ability of the mucus to absorb odorants differentially, could result in some measure of chromatographic separation, possibly leading to characteristic concentration gradients of odorants across the epithelium and variations of concentration with time. This “imposed” spatial patterning has been proposed as a contributory mechanism for odorant recognition (Mozell, 1970; Moulton, 1976; Hornung and Mozell, 1981; Mozell et al., 1987). However, Laing (1988) found no correlation between the absorption properties of an odorant and the human ability to recognize that odorant in mixtures.

B. CONTROL OF SECRETION The control of secretion in the olfactory mucosa has been reviewed by M. L. Getchell et al. (1988). The olfactory mucosa is innervated by the trigeminal nerve. T h e sensory role of this nerve (e.g., Silver et al., 1986) is beyond the scope of this review, but the secretomotor function may play an important modifying role in olfaction. Substance P immunoreactivity is found near the Bowman’s glands and blood vessels of the lamina propria (Papka and Matulionis, 1983); also, fibers extend to near the epithelial surface (Bouvet et al., 1987a; M. L. Getchell et al., 1989; Zielinski et al., 1989a). These fibers terminate in varicosities mainly between the sustentacular cells with no morphological sign of synaptic contact. The fibers are present at a ratio of about one fiber to 30-300 sensory neurons in different amphibian species. Stimulation of the trigeminal nerve or topical application of substance P gives morphological signs of secretion (M. L. Getchell et al., 1989). T h e olfactory mucosa also receives innervation by the terminal nerve (Wirsig and Getchell, 1986) and there are cholinergic terminals between the gland cells of Bowman’s glands and near the adjacent blood vessels (Zielinski et aE., 198913). Vasoactive intestinal peptide may also play a role in secretory control (M. L. Getchell et al., 1987, 1988). There is evidence for adrenergic fibers in the olfactory mucosa (Kawan0 and Margolis, 1985) and for a-adrenergic (Zielinski et al., 1989b) and P-adrenergic (Getchell and Getchell, 1984) regulation of secretion in the olfactory glands. Luteinizing hormone-releasing hormone immunoreactive fibers can be found around the ducts of the anterior medial

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SHIRLEY

glands and close to the blood vessels of the olfactory mucosa in the rat (Zheng ~t al., 1988). Odorants also can induce secretion from the glands of the olfactory mucosa (M. L. Getchell et al., 1987) and this effect can be blocked by a muscarinic cholinergic antagonist, probably indicating a secretomotor reflex. In addition, odorants can stimulate secretion from sustentacular cells in some species (Ekblom et al., 1984; M. L. Getchell et al., 1987), although in man (Moran et al., 1982;Jafek, 1983) and some other species (Yamamoto, 1976) the sustentacular cells seem to be nonsecretory. The control of this process is unclear but it may be relevant that odorants can induce a depolarization of long duration in the supporting cells (Trotier and MacLeod, l986b).

C;. CENTRAL CONTROL OF

THE

SENSORY CELLS

Endocrine processes have an important role in the modulation of olfactory function generally (reviewed in Doty, 1989). This section is concerned only with possible central control of the sensory cells. Application of substance P can evoke a slow electrical potential change from the olfactory tissue (Bouvet et al., 1984), cause changes in the spike activity of receptor neurons (Bouvet ct al., 1988),and modify the responsiveness to odorants (Bouvet et d., 1987b). Stimulation of the trigeminal nerve has the same effects (Bouvet el al., 1987b,c). Substance P immunoreactivity is also found in the olfactory epithelium of fish (Szabo et al., 1987). Acetylcholine also causes slow electrical potentials (Bouvet et al., 1984) and modifies spike activity in the receptors (Bouvet et al., 1988). It seems likely that these events are mediated via receptors on the sensory neurons rather than simply being consequences of changes in mucus secretion. The sensory cells could, therefore, be under some measure of central control. ‘The olfactory nerves d o carry nonodor receptors; peripheral-type benzodiazepine receptors were found by Anholt et al. (1984), and in the sensory cells of the lobster, there is a chloride channel, directly controlled by histamine (McClintock and Ache, 1989a), which seenis to tnodulate spiking activity (Bayer et al., 1989).

I). XENoBIOrIC-METABOLIZIIL’G

ENZYMES

‘Theolfactory mucosa is a rich source of enzymatic activity capable of metabolizing foreign substances. This subject has been reviewed by Dahl (1988)and Dahl et al. (1988). T h e presumed function of these enzymes is the metabolism of odorants to odorless or more soluble compounds,

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although it is possible that they might, by metabolism, change odor quality. These enzymes often have a very wide specificity, and there is no reason to suppose that they do not metabolize endogenous material also. T h e olfactory mucosa contains high levels of cytochrome P-450 (Dahl et al., 1982; Voigt et al., 1985; reviewed by Jenner and Dodd, 1988a), which has different substrate specificities and different kinetic parameters compared to the liver enzyme (Brittebo and Ahlman, 1984; Brittebo and Rafter, 1984). The high levels of activity in the olfactory tissue are due in part to intrinsic differences between the olfactory and liver enzymes (Reed et al., 1988) and in part to enhanced electron flow due to the high ratio of NADPH:cytochrome P-450 reductase:cytochrome P-450 in the olfactory tissue (Reed et al., 1986). Among other functions in this tissue, P-450 has been implicated in the metabolism of steroids (Brittebo and Rafter, 1984; Brittebo, 1982), possibly including exogenous odorous molecules (Gower et al., 1981; Persaud et al., 1988b). The olfactory enzymes and those of other tissues differ in their properties. There are mechanistic differences between olfactory and liver enzymes (Reed and De Matteis, 1989). The olfactory tissue contains unique isoenzymes (Ding and Coon, 1988), different isoenzyme profiles than those found in other tissues (Ding et al., 1986;jenner and Dodd, 1988b; Larsson et al., 1989), and there are differences in inducibility (Ding et al., 1986). Sequence data derived from a cDNA clone (Nef et al., 1989) indicate that an olfactory enzyme, although a member of the P-45011 family, has novel features. Other enzymes that have been found in the olfactory region are FAD-monoxygenase (McNulty and Heck, 1983; McNulty et al., 1983), epoxide hydrolases (Bond, 1983), aldehyde dehydrogenase (CasanovaSchmitz et al., 1984; Bogdanffy et al., 1986), carboxylesterases (Stott and McKenna, 1985; Dahl et al., 1987; Bogdanffy et al., 1987), rhodanase (Dahl, 1989), and the “phase 11” enzymes responsible for transferring water-soluble groups (Bond, 1983; Baron et al., 1986). There are reports of a rather more “odor-specific”metabolizing system in the lobster. T h e adenine nucleotides are important olfactory stimuli for these animals and dephosphorylation enzymes and an adenosine uptake system have been found (Trapido-Rosenthal et al., 1987a,b; Gleeson et al., 1989). E. ODORANT-BINDING PROTEIN

Odorant-binding protein was identified and isolated from olfactory tissue on the basis of its ability to bind pyrazines (Pelosi and Pisanelli, 1981; Pelosi et al., 1982; Bignetti et al. 1985; Pevsner et al., 1985)and was

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originally thought to be a receptor molecule. However, the specificity is weaker than originally believed and the protein will bind many odorants at micromolar concentrations (Topazzini et al., 1985; Pevsner et al., 1986; Bignetti et al., 1988). T h e protein is soluble and exists as a dimer of 2 X 19 kDa (Pevsner et al., 1985). In the cow it is found in the glands of the olfactory and respiratory epithelium (Pevsner et al., 1986) but in the rat it is found in the lateral nasal gland, which discharges just behind the external naris at a considerable distance from the olfactory epithelium (Pevsner et al., 1988a). The protein has been cloned and sequenced (Lee et ul., 1987; Pevsner et al., 1988b) and it is a member of the a2-microglobulin family of carrier proteins. T h e properties of this protein have been reviewed by Snyder et al. (1988b). There have been speculations as to the function of this protein. If an odorant partitions between air and an aqueous medium, it achieves a higher concentration in the liquid than in the gas. Hence, the olfactory mucus does not pose a permeability barrier to odorants and a carrier protein cannot facilitate diffusion under these circumstances; in fact, it will retard diffusion by virtue of its high molecular weight. It is unlikely, therefore, that the odorant-binding protein is part of a simple delivery system, as suggested by Schofield (1988). An alternative suggestion is that the protein-containing secretion is atomized near the naris in the rat to scrub the incoming air and act as part of an odorant delivery system, but the physical design of the olfactory cavity is such that the probability of deposition of an aerosol particle is less than that of deposition of an odorant direct from the gas phase. If the protein serves as a prereceptor or as part of an odor-clearance mechanism for the olfactory epithelium, it is difficult to see why it should be produced so far from the epithelium in the rat and have no mechanism for delivering it there (the flow of mucus is away from the epithelium). Perhaps the most likely function is as a scavenger to keep the air passages in general clean, and it is fortuitous that some of the protein is produced in the olfactory epithelium of the cow.

VII. Conclusion

Much work remains to be done for a full understanding of the olfactory system and it requires collaboration across many different disciplines. Aerodynamics and physical chemistry can help define the concentration profiles of materials in the mucus and lead to an understanding of stimulus dynamics. A great goal for the biochemists and

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molecular biologists is the separation of the various types of receptor molecules, as this leads to an understanding of the response spectrum of each, definitive structure-activity relationships, and the promise of the understanding of the chemistry of odor-receptor interactions. Beyond this it may lead to the ability to “type” cells according to their receptor content, a technique invaluable in the understanding of system function and of the guidance processes involved in regenerative glomerulus selection and synapse formation. The cell physiologists have the problem of the arrangement of the receptor molecules on cells and cellular control, the stoichiometry of the transductory system, and also the mechanisms whereby and the extent to which primary cells act as information processors rather than as simple transducers. T h e system physiologists and neuroanatomists will contribute an understanding of the neural mechanisms involved in olfactory information processing and their changes with learning. T h e fields of psychology, animal behavior, and information theory also have much to contribute. But above all it is necessary to appreciate the relationships between the various fields of study.

References

Admek, G. D., Gesteland, R. C., Mair, R. C., and Oakley, B. (1984). Bruin Res. 310, 87-97. Akerson, R. A. (1988). I n “Molecular Neurobiology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 297-318. Plenum, New York. Allen, W. K., and Akerson, R. (1985a).J. Neurosci. 5, 284-296. Allen, W. K., and Akerson R. (1985b). Dev. Bid. 109, 393-401. Alonso, J. R., CoveAas, R., Lara, J., De Leon, M., and A@n, J. (1 989). Bruin Res. 490,385390. Amoore, J. E. (1970). “The Molecular Basis of Odor.” Thomas, Springfield, Illinois. Anderson, P. A. V., and Ache, B. W. (1985). Bruin Res. 335, 273-280. Anderson, P. A. V., and Hamilton, K. A. (1987). Neuroscience 21, 167-173. Anholt, R. R. H. (1987). Trends Biochem. Sci. 12, 58-62. Anholt, R. R. H. (1988). BiochemGtry 27, 6464-6469. Anholt, R. R. H., Murphy, K. M. M., Malk, G., and Snyder, S. H. (1984).J. Neurosci. 4,593603. Anholt, R. R. H., Aebi, U., and Snyder, S. H. (1986).J. Neurosci. 6, 1962-1969. Anholt, R. R. H., Mumby, S. M., Stoffers, D. A., Girard, P. G., Kuo, J. F., Gilman, A. G., and Snyder, S. H. (1987a). Ann. N.Y. Acud. Sci. 510, 152-156. Anholt, R. R. H., Mumby, S. M., Stoffers, D. A., Girard, P. G., Kuo, J. F., and Snyder, S. H. (1987b), Biochemistry 26, 788-795. Asanuma, N., and Nomura, H. (1989), Chem. Senses 14, 323. Astic, L., and Saucier, D. (1986). Bruin Res. Bull. 16, 445-454. Astic, L., and Saucier, D. (1988). Dev. Brain Res. 42, 297-304. Astic, L., Saucier, D., and Holley, A. (1987). Brain Res. 424, 144-152.

42

S. G. SHIRLEY

Astic, L., Saucier, D., Jourdan, F., and Holley, A. (1988).Chem. Senses 13, 333-344. Astic, L., Le Pendu, J., Mollicone, R., Saucier, D., and Oriol, R. (1989).J. Comp. Neurol. 289, 386-394. Aston-Jones, G. (1985). Phyzol. Psychol. 13, 1 18-126. Baker, H. (1986a),Exp. Bruin Rps. 65, 245-249. Baker, H. (1986b). J . Comp. Nmrol. 252, 206-226. Baker, H. (1988). Irt “Molecular Biology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 185-216. Plenum, New York. Baker, H., Grillo, M..and Margolis, F. L. (1989).J. Comp. ‘Veurol. 285, 246-261. Barber, P. C. (1982a). rVeurosczerLce7, 2677-2685. Barber, P. C.(198%). Bib/. Arml. 23, 12-25. Barber, P. C. (1989). zVeurosctPtire 30, 1-9. Barher, P. C.,and Ddhl, D. (1987). Esp. Bruit1 Ke.s. 65, 681-685. Barber, P. G., a n d Jensen, S. (1988). I n “Molecular Neurobiology of the Olfactory System: Molecular. Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, etis.), p p 333-352. Plenum, New York. Barber, P. C., and Lindsay, R. M. (1982). Neuro.scie~~ce 7 , 3077-3090. Baron, J., Voigt, J. M., Whitter, T. B., Kawabata, T., Knapp, S. A , , Guengerich, F. P., and Jacoby, W. B. ( 1986). Itr “Biological Reactive intermediates. 111. Molecular and Celiui;rr Mechanisms of Action in Animal Models and Human Disease” (R. Snyder, ed.), pp. 109-141. Plenum, New York. Baskin, D. G.. Wilrox, B. J . , Figlewicz, D. P.. and Dorsa, D. M. (1988). T r e r d Neurosci. 11, 107-1 1 1 . Bayer. I-.A., McClinttxk, T. S., Griinert, U . , and Ache, B. W. (I989).J. Exp. Biol. 145, 133146.

Bell, G. A., Laing. D. G., and pdnhuber, H . (1987). Brain Res. 426, 8-18. Benson, T. E.. B u d , G. D., Greer, C . A., Landis, D. M. D., and Shepherd, G. M. (1985). Braiir Res. 339, 67-68. Bignetti, E., Cavaggioni, A., Pelosi, P., Persaud, K. C.,Sorbi, K. T., and Tirindelli, R. (1985). Eur. J . Bzorhem. 149, 227-231. Bignetti. E.. Cattaneo. P., Cavaggictni, A., Damiana, G., and Tirindelli, R. (1988).Comp. B?UC/letti, P/LJ,>iOl.90, 1-7. Bogdanffy, M. S., Randall, N.W., and Morgan, K. T. (1986). Toxicol. Appl. Pharmacol. 82, 560-567. Bogdanffy, ,M. S., Randall, H. W., and Morgan, K. T. (1987). Toxirol. Appl. Phrmacol. 88, 183- 194. Bond, J . A. (1983). Currcer KPJ. 43, 4804-481 1. Borroni, P. F., Handrich, L. S., and Atema, J. (1986). Behav. Neurosri. 100, 206-212. Bouvet, J. F., Delaleu, J. C., and Hoiley, A. (1984). C.R. Hebd. Seritzce.s A d . Sci. 298, 169172. Bouver, J. F., Godinor, F., Croze, S., and Delaleu, J. C. (l987a). Cherrt. Sences 12, 499-506. Boiivet, J. F., Delaleu. J. C.,and Holley, A. (1987b). Anrc. N.Y. Acad. Sci. 510, 187-189. Bouvet, J. F.. Delaleu. J. C., and Holley, A. (1987~). ~ V p u m c iLett. . 7 7 , 181-187. Uouvet, J. F.. Delaleu, J . C.,a n d Holley, A. (1988). Neurosci. Kes. 5 , 214-223. Boyle, A. G.. Park, Y. S.. Huque, T., and Bruch, R. C. (1987). Conip. Biochem. Physiol. H 88B, 767-775. Breipohl, U:.and Fernandez, M. (1977). Cell Tissue Res. 183, 105-1 14. Breipohl. W.. Bijvank, G. J.. and Zipple, H. P. (1973).Z. Zell/on,srh.M i k r o ~ kA7mt. . 138,4394.54.

OLFACTION

43

Breipohl, W., Mackay-Sim, A., Grandt, D., Rehn, B., and Darrelmann, C. (1986). In “Ontogeny of Olfaction” (W. Breipohl, ed.), pp. 21-34. Springer-Verlag, Berlin. Brittebo, E. B. (1982).Actu Phunnacol. Toxicol. 51, 441-445. Brittebo, E. B., and Ahlman, M. (1984). Chem.-Biol. Interact. 50, 233-245. Brittebo, E. B., and Rafter, J. (1984).J. SteroidBiochem. 20, 1147-1151. Brown, D., Garcia-Segura, L.-M., and Orci, L. (1984). Histochemistq 80, 307-309. Brown, S. B., and Hara, T. J. (1981). Biochim. Biophys. Acta 675, 149-162. Brown, S. B., and Hara, T. J. (1982). In “Chemoreception in fishes” (T.J. Hara, ed.), pp. 159-180. Elsevier, New York. Bruch, R. C., and Huque, T. (1987). Ann. N.Y. Acud. Sci. 510,205-207. Bruch, R. C., and Kalinoski, D. L. (1987).J. Biol. Chem. 262, 2401-2404. Bruch, R. C., and Rulli, R. D. (1988). Comp. Biochem. Physiol. B 91B, 533-540. Bruch, R. C., Kalinoski, D. L., and Kare, M. R. (1988). Ann. Rev. Nutr. 8, 21-42. Brunjes, P., and Frazier, L. L. (1986). Bruin Res. Rev. 11, 1-45. Burd, G. D., Davis, B. J., Macrides, F., Grillo, A., and Margolis, F. L. (1982).J. Neurosci. 2, 244-255. Cagan, R. H., and Zeiger, W. N. (1978). Proc. Nutl. Acud. Sci. U.S.A. 75, 4679-4683. Cancalon, P. (1978). Chem. Senses Flavour 3, 381-396. Caprio, J., Dudek, J., and Robinson, J. J., I1 (1989).J. Cen. Physiol. 93, 245-262. Carr, V. M., Farbman, A. I., Lidow, M. S., Colletti, L. M., Hempstead, J. L., and Morgan, J. I. (1989).J. Neurosci. 9, 1179-1198. Carr, W. E. S. (1987). In “Sensory Biology of Aquatic Animals” (J. Atema, A. N. Popper, R. R. Fay, and W. N. Tavolga, eds.), pp. 3-28. Springer-Verlag, Berlin. Carr, W. E. S., and Derby, C. D. (1986a).J. Chem. Ecol. 12,989-101 1. Carr, W. E. S., and Derby, C. D. (198613). Chem. Senses 11, 49-64. Carr, W. E. S., Gleeson, R. A., Ache, B. W., and Milstead, M. L. (1986).J.Comp. Physiol. 158, 33 1-338. Carr, W. E. S., Gleeson, R. A., Ache, B. W., and Milstead, M. L. (1987).Ann. N.Y. Acud. Sci. 510, 219-221. Casanova-Schmitz, M., David, R. M., and Heck, H. D. (1984). Biochem. Phurmacol. 33, 1137-1 142. Chaput, M. A. (1986). Physiol. Behuu. 36, 319-324. Chaput, M. A., and Holley, A. (1985). Physiol. Behuu. 34, 249-258. Chaput, M. A., and Lankeet, M. J. (1987). Physiol. Behau. 40, 453-462. Chen, Z., and Lancet, D. (1984). Proc. Nutl. Acad. Sci. U.S.A. 81, 1859-1863. Chen, Z., Pace, U., Heldman, J., Shapira, A., and Lancet, D. (1986a).J. Neurosci. 6, 21462154. Chen, Z., Pace, U., Ronen, D., and Lancet, D. (1986b).J. Biol. Chem. 261, 1299-1305. Chiflikian, M. D., Kilmin, M., Galoyan, A. A., and Hajos, F. (1986). Neurochem. Res. 11, 1597-1608. Coon, H. G., Curcio, F., Sakaguchi, K., Drandi, M. L., and Swerdlow, R. D. (1989). Proc. Natl. Acud. Sci. U.S.A. 86, 1703-1708. Coopersmith, R., and Leon, M. (1984). Science 225,849-851. Coopersmith, R., and Leon, M. (1986). Bruin Res. 371, 400-403. Coopersmith, R., and Leon, M. (1989).J . Comp. Neurol. 289, 348-350. Coopersmith, R., Henderson, S. R., and Leon, M. (1986). Deu. Bruin Res. 27, 191-197. Costanzo, R. M., and Morrison, E. E. (1989).J. Neurocytol. 18, 381-391. Costanzo, R. M., and Mozell, M. M. G. (1976).J. Gen. Physiol. 68, 297-312. Costanzo, R. M., and OConnel, R. J. (198O).J. Cen. Physiol. 76, 53-68. Cuschieri, A,, and Bannister, L. H. (1975).J.Anut. 119, 277-286.

44

S. G.

SHIRLEY

Dahl. A. R. (1988).I n “Molecular Neurobiology o f t h e Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 51-70. Plenun~,New l’ork. Dahl, A. R. (1989). Trxicol. Lrtl. 45, 199-206. Dahl, A. R., Hadle); W‘.M., H a m , F. F., Benson, J. M.,and McClellan, R. 0. (1982). Science 216, 57-59. Dahl, A . R . , Miller, S. C., and Petridou-Fischer, J. (1987). To.rUo1. Lett. 36, 129-136. Dahl,A . K., Bond, J. M..Petridou-Fischer, J., Sabourin, P. J., and Whaley, S. J . (1988). Toxirot. App1. Phaminro!. 93, 484-492. Daval, (;., and Leveteau, J. (1982). C.K. Hrbd. Smnres Acad. Sci. 295, 637-640. Dawson, V. L., Dawson, T. M., and M’amsley, J. K. (1988). Irr “h$olecular Neurobiology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 99-1 17. Plenum, New York. Delaleu, J. C., and Holley, A. (1980). Chmi. Srturs Flavour 5, 20.5-218. Delaieu, J. C., and Holley. A. (1983). Nrurosci. Lett. 37, 251-256. Denis-Donini, S. (1989).,\‘citurr (London) 339, 701-703. Derby, C. D., a n d Ache, B. W’. (1984a). Chew. Sexses 9, 201-218. Derbv, C. D., and Ache, B. W. (1984b).J. A‘riirophysiol. 51, 906-924. Derby, C. D., Carr, b‘.E. S., and Ache, B. W’. (1984).J. Comp. Pliy.siysiol. A 155A, 341-349. Derby, C. D., Ache, B. %’., and Kennel, E. W. (1985). Chrrri. SCJISPS10, 301-316. Derby, C. D., Carr, iV. E. S., and Ache, B. W’.(1987). A m . N.Y. Acad Sci. 510, 250-253. DeSimone, J. A,, Persaud, K. C., and Heck, G. L. (1988).I I I “Molecular Neurobiology of the Olfactor) System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 159-181. Plenum, New York. Dickinson, C., and Keverne, E. B. (1988). Phjsiosiol. Behail. 43, 313-316. Dickinson, K. (1987). Ph.D. Thesis, University of M‘arwick. Ding, X.,and Coon, M.J. (1988).Biorhrniirfq 27, 8330-8337. Ding, X.,Koop, D. R.. Crump, B. L., and Coon, M. J . (1986).Xlol. Phnrwiacol. 30,370-378. Dionne, \I. E. (1987). A w . S . Y . Acnd. Sci. 510, 258-259. . 34, 143-156. Doroslienko. hl. A., and Motavkin, P. A. (1986). Arta M o r ~ h o lHung. Doty, K.L. (I9X8). Exprrientio 42, 257-271. Doty, R. L. (1989). I n “Keur-a1 Control of Reproductive Function” 0. M. Lakoski, J. R. Perez, and D. K. Kassin, eds.), pp. 567-582. Liss, New York. Deving, K. B. ( 1 987). Acfa Physiol. Scand. 130, 285-298. Duchainp, A. (3982). Chon. Snnr.5 7, 191-210. Duchanip, A., and Sicard, G. (1984). Chrrri. Senses 8, 335-366. Duchamp-\’iret, P., Duchamp, A , , and Vigouroux, M. ( 1989). J . ”Vurophysiol. 61, 1085-1094. Edwards, D. A , Mather, R. A,, and Dodd. G. H. (1988). Exprrirnticl 44,208-21 1. Ekbliini. A.. Flock. A., Hansson, P., and Ottoson, D. (1984). cic/n O f o - L ~ q ~ ~ 98,35 g o l . 1-361. EngstrOtn, B . , Ekbioni. A., a n d Hanssoii, P. (1989). Otola~iiy~igologica 108, 259-267. Farbmau. A. I. ( 1988). In “%folecular Neurobiology of the Olfactory System: Mokcular, Membranous a n d Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 319-332. Plenutn, New York. Farlman, A. I., and Menco, B. P. M. (1986). I i i “Ontogeny of Olfaction” ( W Breipohl, ed.), pp. 4.3-.36. Springer-Verlag, Berlin. Fescnko. E. E.. Novoselov, V. I . , and Bystrova, M. F. (1987). FEBS Lrtt, 219, 224-227. Feseiiko, E. E., N o v ~ ~ V.KI.,~ and ~ v Bystrova, , hl. F. (1988).Biochini. BiopltyJ. A d a 937,36937x. Firestein, S., and M’erblin, F. S. (1987a). Pror. Nall. Accld. Sci. U.S.A. 84, 6292-6296.

OLFACTION

45

Firestein, S., and Werblin, F. S. (1987b). Ann. N.Y. Acad. Sci. 510, 287-289. Firestein, S.,and Werblin, F. S. (1989). Science 244, 79-82. Freeman, W. J., and Baird, B. (1987). Behav. Neurosci. 101, 393-408. Freeman, W. J., and Grajski, R. (1987). Behav. Neurosci. 101, 766-777. Freeman, W. J., and Schneider, W. (1982). Psychophysiology 19,44-56. Frings, S.,and Lindermann, B. (1988).J. Membr. Biol. 105,233-243. Fujita, S. C., Mori, K., Imamura, K., and Obata, K. (1985). Brain Res. 326, 192-196. Fuzessery, Z.M., Carr, W. E. S., and Ache, B. W. (1978). Biol. Bull. (Woods Hole, Mass.) 154, 226-240. Gervais, R. (1987). Brain Res. 400, 151-155. Gervais, R., Holley, A., and Keverne, B. (1988). Chem. Senses 13, 3-12. Gesteland, R. C. (1975).In “Methods in Olfactory Research” (D. G. Moulton, A. Turk, and J. W. Johnson, Jr., eds.), pp. 269-323. Academic Press, London. Gesteland, R. C., Yancey, R. A., and Farbman, A. I. (1982). Neuroscience 7, 3127-3136. Getchell, M. L., and Gesteland, R. C. (1972). Proc. Natl. Acad. Sci. U.S.A. 69, 1494-1498. Getchell, M. L., and Getchell, T. V. (1984).J. Comp. Physiol. A 155A,435-443. Getchell, M. L., Zielinski, B., DeSimone, J. A., and Getchell, T. V. (1987).J. Comp. Physiol A 160A, 155-168. Getchell, M. L., Zielinski, B., and Getchell, T. V. (1988). In “Molecular Neurobiology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 71-98. Plenum, New York. Getchell, M. L., Bouvet, J. F., Finger, T. E., Holley, A., and Getchell, T. V. (1989). Cell Tissue Res. 256, 381-390. Getchell, T. V. (1974).J. Neurophysiol. 37, 1115-1 130. Getchell, T. V. (1986). Physiol. Rev. 66, 772-817. Getchell, T. V. (1988). Neurosci. Lett. 91, 217-221. Getchell, T. V., and Shepherd, G. M. (1978).J. Physiol. (London) 282, 541. Getchell, T. V., Heck, G. L., DeSimone, J. A., and Price, S. (1980). Bi0phys.J. 29, 397-412. Getchell, T.V., Margolis, F. L., and Getchell, M. L. (1984). Prog. Neurobiol. 23, 317-345. Gibson, T. R., Zyskind, A. D., and Glembotski, C. C. (1988). J. Neurosci. 8, 3067-3074. Gilman, A. G. (1987). Annu. Rev. Biochem. 56, 615-649. Giradot, M. N., and Derby, C. D. (1988).J. Neurophysiol. 60, 303-324. Gladysheva, O.,Kukushkina, D., and Martynova, G. (1986). Acta Histochem. 78, 141-146. Gleeson, R. A,, and Ache, B. W. (1985). Brain Res. 335, 99-107. Gleeson, R. A,, Trapido-Rosenthal, H. G., Littleton, J. T., and Carr, W. E. S. (1989). Comp. Biochem. Physiol. C 92C,413-418. Glembotski, C. C., Wildey, G. M., and Gibson, T. R. (1989). Cell. Mol. Neurobiol. 9, 57-74. Gold, G.H., and Nakamura, T. (1987). Trends Pharmacol. Sci. 8, 312-316. Goldberg, S. J., Turpin, J., and Price, S. (1979). Chem Senses 4, 207-214. Gonzales-Estrada, M. T., and Freeman, W. J. (1980). Brain Res. 202, 373-386. Cower, D. B., Hancock, M. R.,and Bannister, L. H. (1981). In “Biochemistry of Taste and Olfaction” (R.H. Cagan and M. R. Kare, eds.), pp. 8-28. Academic Press, New York. Gray, C. M., Freeman, W. J., and Skinner, S. R. (1986). Behuv. Neurosci. 100, 585-596. Graziadei, P. P. C., and Monti-Graziadei, G. A. (1986a). Ann. N.Y. Acad. Sci. 457, 127-142. Graziadei, P. P. C., and Monti-Graziadei, G. A. (198613).Neuroscience 19, 1025-1035. Gross-Isseroff, R., and Lancet, D. (1988). Chem. Senses 13, 191-204. Halftsz, N., and Shepherd, G. M. (1983). Neuroscience 10, 579-619. Hamilton, K. A., and Kauer, J. S. (1985). Bruin Res. 338, 181-185. Hamilton, K. A,, and Kauer, J. S. (1987). Ann. N.Y. Acad. Sci. 510, 332-334. Hamilton, K. A., and Kauer, J. S. (1988).J. Neurophysiol. 59, 1736-1755.

46

S. G . SHIRLEY

Hamilton, K. A., and Kauer, J. S. (1989).J. ,Veurophysiol. 62, 609-625. Handrich, L. S.. and Atema, J. (1987). Ann. M.Y. Acad Sci. 510, 342-344. Harrison, ‘I.A.. and Scott, J. W. (1986).J. Neuruphysiol. 56, 1571-1589. Hedlund, B.. Masukawa, L. M., and Shepherd, G. M. (1987).J. Neurosci. 7, 2338-2343. Heldnian. J., and Lancet, D. (1986).J. Neurochem. 47, 1527-1533. Hinds, J. W., Hinds, P. L., and McNelly, N. A. (1984). A m t . Rec. 210, 375-383. Hokin. L. E. (1985). Annu. Rat. Biochem. 54, 205-235. Hornung, D. E., and Mozell, M. M.(1981). In “Biochemistry of Taste and Olfaction” (R. H . Cagan and M. R. Kare, eds.), pp. 33-45. Academic Press, New York. Hornung, D. E., Youngentob, S. L.. and Mozell. M. M. (1987). Brain Res. 413, 147-155. Houslay. M.D., and Gordon, L. XI. (1983). Curr. Top. Membr. Tramp. 18, 179-231. Huque, %, and Bruch, R. C . (1986). Biorhem. B1ophy.c. Res. Commun. 137, 36-42. .Jafek, B. M’. (1983). Laygoscope 93, 1576-1599. Jastrelmotf, P. J., Penderson, P. E., Greer, C . A., Stewart, W. B., Kauer, J. S., Benson, T. E., and Shepherd, G. M.(1984). Proc. Natl. Acad. Sci. U.S.A. 81, 5250-5254. .Jenner, ,J.. and Dodd, G . H. (1988a). Drug Metab. Drug Interact. 6, 123-148. Jenner, J., and Dodd. G . H. (1988h). Bzorhem. Phannacol. 37, 558-559. Jiang. T.. and Holle); A. (1987). Anti. N.Y. Arad. Sci. 510, 384-387. Johnson, B. K.,and Atema,.J. (1983). Neurosri. Lett. 41, 145-150. Johnson, B. R.. Borroni, P. F., and Atema, J. (1985). Cheni. Senses 10, 367-373. Johnson, B. R., Merrill, C. I., Ogle, K. C., and Atema, J. (1987). Ann. N.Y. Acad. Sci. 510, 388-590. Johnson, B. R., Merrill, C. L., Ogle, K. C., and Atema, J. (1988a).J. Comp. Physiol. A 162A, 201 -2 12. Johnson, K. R., Farbnian, A. I., and Gonzales, F. (1988b).J. Neurobiol. 19, 681-694. Jones, D. ‘I..and Reed, K. K. (1987).J. Biol. Clmz. 262, 14241-14250. Jones, D. T., a n d Reed, R. R. (1989). Science 244, 790-795. Jones, D.T.. Barbosa, E., and Reed, R. R. (1988). Cold Spring Harbor Symp. @ant. Bzol. 53(Pt. I), 349-3.54. Joshi, H., Getchell, M. L., Zielinski, B., and Getchell, T. V. (1987). Neurosci. Lett. 82,321326. Jourdan, F.. Duveau, A., Astic, L., and Holley, A. (1980). Brain Res. 188, 139-154. Kaba, H.. and Keverne, B. (1988). Neuroscience 25, 1007-1012. Kaba, H . , Rosser, A., and Keverne, B. (1989). h‘eurosczence 32, 657-662. KaOta, W. A. (1987).J. Comp. Physiol. A 161A, 867-880. Kaissling, K.-E. (1986). Annu. Rev. Neurosci. 9, 121-145. Kaissling, K.-E. (1987). I n “K.H. Wright Lectures on Insect Olfaction (K. Colbow, ed.), pp. 12 1-1 39. Simon Fraser University, Burnaby, B.C., Canada. Kalinoski. D. L . , Bruch. R. C.,and Brand, J. G. (1987). Brain Res. 418, 34-40. Karpov, A. P. (1980). In “Neural Mechanisms of Goal Directed Behavior and Learning” (R. F. ‘Thompson, L. H . Licks, and V. R. Shyrkov, eds.), pp. 273-282. Academic Press, New York. Krshiwayanagi, %I., and Kurihara, K. (1984). Brain Re.?. 359, 97-103. Kashiwayanagi, M., Sai, K., and Kurihara, K. (1987a). Anti. N.Y. Acad. Sci. 510, 398-399. Kashiwayanagi, M., Sai, K., arid Kurihara, K. (1987b). J. Gem Physiol. 89, 443-459. Kashixcayanagi, M., Shoti, T., and Kurihara, K. (1988). Biorheni. Biophys. Res. Commun. 154, 497-442. Katawit, W., Nakao, K.. Morii, N.,Kiso, Y., Yaniashita, H.. Iniura, H., and Sano, Y. (1986). Stwrosciencr 16, 52 1-546. Kaiier, J. S. (1974). J . Physiol. ( L o n d ~ n243, ) 695-7 15.

OLFACTION

47

Kauer, J. S. (1980). In “Olfaction and Taste VII” (H. Van der Starre, ed.), pp. 227-236. IRL Press, London. Kauer, J. S. (1981). Anat. Rec. 200, 331-336. Kauer, J. S. (1987). In “Neurobiology of Taste and Smell” (T. E. Finger and W. L. Silver, eds.), pp. 205-231. Wiley, New York. Kauer, J. S. (1988). Nature (London) 331, 166-167. Kauer, J. S., and Hamilton, K. A. (1987). Ann. N.Y. Acud. Sci. 510,400-402. Kauer, J. S., and Moulton, D. G. (1974).J. Physiol. (London) 243, 717-737. Kauer, J. S., Sensemann, D. M., and Cohen, L. B. (1987). Bruin Res. 418, 255-261. Kawano, T., and Margolis, F. L. (1985). Chem. Senses 10, 353-356. Key, B., and Giorgi, P. P. (1986). Neuroscience 18, 507-515. Klein, S. L., and Graziadei, P. P. (1983).J. Comp. Neurol. 217, 17-30. Kosaka, T., Kosaka, K., Heizmann, C. W., Nagatsu, I., Wu, J.-Y., Yanihara, N., and Hama, K. (1987). Bruin Res. 411, 373-378. Kropf, R., Lancet, D., and Lazard, D. (1987). SOC.Neurosci. Abst. 13, 1410. Kubie, J., Mackay-Sim, A., and Moulton, D. G. (1980). In “Olfaction and Taste VII” (H. van der Starre, ed.), pp. 163-166. IRL Press, London. Kurahashi, T. (1989).J. Physiol. (London) 419, 177-192. Kurahashi, T., and Shibua, T. (1989). Chem. Senses 14, 323. Kurihara, K., and Koyama, N. (1972). Biochem. Biophys. Res. Comrnun. 48, 30-34. Kurihara, K., and Yoshii, K. (1983). Bruin Res. 274, 239-248. Labarch, P., and Bacigalupo, J. (1988).J. Bioenerg. Biomembr. 20, 551-570. Labarch, P., Simon, S. A., and Anholt, R. R. H. (1988). Proc. Nutl. Acud. Sci. U.S.A. 85,944948. Laffort, P., and Gortan, C. (1987). Chem. Senses 12, 139-142. Laing, D. G. (1986). Physiol. Behav. 37, 163-170. Laing, D. G. (1988). Chem. Senses 13, 463-472. Lancet, D. (1986). Annu. Rev. Neurosci. 9, 329-355. Lancet, D. (1988).In “Molecular Biology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 25-50. Plenum, New York. Lancet, D., and Pace, U. (1987). Trends Biochem. Sci. 12, 63-67. Lancet, D., Greer, C. A., Kauer, J. S., and Shepherd, G. M. (1982). Proc. Nutl. Acud. Sci. U.S.A. 79, 670-674. Lancet, D., Chen, Z., Ciobotariu, A., Eckstein, F., Khen, M., Heldman, J., Ophir, D., Shafir, I., and Pace, U. (1987). Ann. N.Y. Acud. Sci. 510, 27-32. Lancet, D., Lazard, D., Heldman, J., Khen, M., and Nef, P. (1988). Cold Spring Harbor Symp. Quant. Biol. 53(Pt. I), 343-348. Larsson, P., Pettersson, H., and Tjalve, H. (1989). Curcinogenesis (London) 10, 11 131118.

Lazard, D., Barak, Y., and Lancet, D. (1989). Biochim. Biophys. Actu 1013, 68-73. Lee, K. H., Wells, R. G., and Reed, R. R. (1987). Science 235, 1053-1056. Leon, M. (1987). Trends Neurosci. 10, 434-438. Lerner, M. R., Reagan, J., Gyorgyi, T., and Roby, A. (1988). Proc. Nutl. Acud. Sci. U.S.A. 85, 261-265. Leveteau, J., Andriason, I., Trotier, D., and MacLeod, P. (1989). Chem. S m e s 14,611-620. Lidow, M. S., Gesteland, R. C., Kleene, S. J., and Shipley, M. T. (1987a).Ann. N.Y. Acad. Sci. 510,454-455. Lidow, M. S., Gesteland, R. C., Shipley, M. T., and Kleene, S. J. (1987b). Dev. Bruin Res. 31, 243-258.

48

S. G. SHIRLEY

Lowe, G., Nakamura, T., and Gold, C. H . (1989).Proc. ivatl. Acad. Sci. U.S.A. 86, 56415646. Lynch, J . W., and Barry, P. H. (1989). Biophjs. J. 55, 755-768. Mackav-Sim, A., and Kubie, J. L. (1981). Chem. Senses 6, 249-257. Mackay-Sim, A., and Nathan, M. H. (1984). Anat. Embryol. 170, 93-98. Mackay-Sim, A., and Shaman, P. (1984). Brain Res. 297, 207-216. Mackay-Sim, A., Shaman, P., and Moulton, D. G. (1982). J. Neurophysiol. 48, 584-596. Mackay-Sim, A., Breipohl, W., and Kremer, M. (1988). Exp. Bruin Res. 71, 189-198. Macrides, F., and Davis, B. (1983). In “Chemical Neuroanatomy” (P. C . Emerson, ed.), pp. 391-426. Raven Press, New York. Macrides, F., Schoenfeld, T. A, Marchand, J. E., and Clancey, A. N. (1985).Chem. Senses 10, 175-202. Mair, R. G. (1982a). J . Phjsiol. (Londm) 326, 341-359. Mair, R. G. (1982b). J. Phjsiol. (London) 326, 361-369. . ( 1988). Expeiieiitia 42, 2 13-223. ., and (;esteland, R. C . (1982). Neuroscienre 7, 3117-3125. Mair, R. G.. Gesteland, R. C., and Blank, D. L. (1982). Neuroscience 7, 3091-3103. Margolis. F. L. (1972). Proc. Natl. A c d . Sci. U.S.A. 69, 1221-1224. Margolis, F. L. (1988).In ”Molecular Neurobiology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 237-26.5. Plenum, New York. Margolis, F. L., Grillo, M., Kawano, T., and Farbman, A. I. (1985).J.Neurnchem. 44, 14591464. Margolis, F. L., Grillo, hi., Hempstead, J., and Morgan, J. I. (1987).J. Neurochem. 48,293300. Mason, J. R., Leong. F.-C., Plaxco, K. W., and Morton, T. H. (1985).J. Am. Chem. Soc. 107, 6075-6084. Mason, J. R., Johri, K. K., and Morton, T. H. (1987a).J. Chem. Ecol. 13, 1-18. Mason, j. K.,CLark, L., and Morton, T. H. (1987b). Ann. N.Y. A c d . Sci. 510, 468-471. Masukawa. L. M., Hedlund, B., and Shepherd, G. M. (l985).J. Neurosci. 5, 136-141. Masukawa, L. hl., Hedlund, B., and Shepherd, G. M. (1987).Ann. N.Y. Acad. Sci. 510,475477. Mathews, D. F. (1972). Brain Re.,. 47, 389-400. Matsumoto, H.. and Rhoads, D. E. (199O).J.Neurochpm. 54, 347-350. Matsutani, S., Senba, E., and Tohyama, M. (1988).J. Comp. Neurol. 272, 331-342. Matsutani, S., Senba, E., and Tohyama, M. (1989a). J. Conrp. Neurol. 285, 73-82. Matsutani, S., Senba, E., and Tohyama, M. (1989b).J. Comnp. Neurol. 280, 577-587. Maue, R. A,, and Dionne, V. E. (1987a). pfiuegers Arrh. 409, 244-250. Maue, R. A, and Dionne, V. E. (1987b).J. Gen. Physiol. 90, 95-126. Maue, R. A.. and Dionne, V. E. (1988). In “Molecular Neurobiology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 143- 158. Plenum, New York. McClintock. T. S.. and Ache, B. W. (1989a). Proc. Natl. Arad. Sci. U.S.A. 86, 8137-8142. McClintock, T. S., and Ache, B. \V. (1989b). Chem. Senses 14, 637-648. McXultv, M. J.. and Heck, H. D. (1983). Drug Metab. Dispos. 11, 417-420. McNultv, M. J., (;asanova-Shmitz, M., and Heck, H. D. (1983). Drug Metah. 0i~po.r.11, 421-425. McQueen, J. K.. Martin, M. 1.. and Fink, C. (1988).Neuroendocrinology 47, 437-444. Menco, B. P. M. (1977). Cumtnuia. Agru. Unzv. Wugmiizgmz 77-13, 1-157. Menco, B. P. M. (1980a). Cell Tissue Res. 207, 183-209. Menco, B. P. M. ( 1980b). Cell Ti~rueReg. 21 1, 5-29.

OLFACTION

49

Menco, B. P. M. (1980~).Cell Tissue Res. 211, 361-373. Menco, B. P. M. (1980d). Cell Tissue Res. 212, 1-16. Menco, B. P. M. (1983). I n “Nasal Tumors in Animals and Man” (G. Reznik and S. F. Stinson, eds.), Vol. 1, pp. 45-102. CRC Press, Boca Raton, Florida. Menco, B. P. M. (1985).J. Cell Sci. 78, 31 1-336. Menco, B. P. M. (1988a). Anat. Embryol. 178, 309-326. Menco, B. P. M. (1988b). Anat. Embryol. 178, 381-388. Menco, B. P. M. (1989a). Cell Tissue Res. 256, 275-281. Menco, B. P. M. (1989b). Scanning Microsc. 3, 257-272. Menco, B. P. M., and Farbman, A. I. (1985).J. Cell Sci. 78, 283-310. Menco, B. P. M., Dodd, G. H., Davey, M., and Bannister, L. H. (1976).Nature (London) 263, 597-599. Menevse, A,, Dodd, G. H., and Poynder, T. M. (1977). Biochem. Biophys. Res. Commun. 77, 671-677. Menevse, A., Dodd, G., and Poynder, T. M. (1978). Bi0chem.J. 176, 845-854. Meredith, M. (1986).J. Neurophysiol. 56, 572-597. Meredith, M., and Moulton, D. G. (1978).J. Gen. Physiol. 71, 615-643. Minor, A. V., and Sakina, N. L. (1973). Neirojiziologiya 5, 415-422. Miragall, F., Kadmon, G., Husrnann, M., and Schachner, M. (1988). Dev. Biol. 129, 516531. Mollicone, R., Trojan, J., and Oriol, R. (1985). Dev. Brain Res. 17, 275-279. Monod, B., Mouly, A. M., Vigouroux, M., and Holley, A. (1989).Behav. Brain Res. 33, 5163. Monti-Graziadei, A. G., and Graziadei, P. P. C. (1989). Brain Res. 484, 157-167. Monti-Graziadei, A. G., and Morrison, E. E. (1988). Brain Res. 455, 401-406. Moran, D. T., Rowley, J. C., 111, Jafek, B. W., and Lovell, M. A. (1982).J. Neurocytol. 11, 721-746. Mori, K. (1987a). Prog. Neurobiol. 29, 275-320. Mori, K. (1987b). Brain Res. 408, 215-221. Mori, K., Fujita, S. C., Imamura, K., and Obata, K. (1985).J. Comp. Neurol. 242, 214-229. Morrison, E. E., and Costanzo, R. M. (1989).J. Neurocytol. 18, 393-405. Moulton, D. G. (1976). Physiol. Rev. 56, 578-593. Mozell, M. M. (1970).J. Gen. Physiol. 56, 46-63. Mozell, M. M., Sheehe, P. R., Hornung, D. E., Kent, P. F., Youngentod, S. L., and Murphy, S. J. (1987). J. Gen. Physiol. 90, 625-650. Murphy, R. B. (1988). I n “Molecular Neurobiology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. Getchell, eds.), pp. 121-142. Plenum, New York. Nakamura, T., and Gold, G. H. (1987). Nature (London) 325, 442-445. Nef, P., Heldman, J., Lazard, D., Margalit, T., Jaye, M., Hanukoglu, I., and Lancet, D. (1989).J. Biol. Chem. 264, 6780-6785. Neidle, A., and Kandera, J. (1974). Brain Res. 80, 359-364. Neuhaus, W. (1981). Z. Saugetier Kd. 46, 301-310. Nomura, T., and Kurihara, K. (1987a). Biochemistry 26, 6135-6140. Nomura, T., and Kurihara, K. (1987b). Biochemistry 26, 6141-6144. Nomura, T., and Kurihara, K. (1989). Biochim. Biophys. Acta 1005, 260-264. Novoselov, V. I., Krapivinskaya, L. D., and Fesenko, E. E. (1988a). Chem. Senses 13, 267278. Novoselov, V. I,, Krapivinskaya, L. D., Krapivinsky, G. B., and Fesenko, E. E. (1988b). FEBS Lett. 234,471-474. OConnel, R.J. (1986). Experientia 42, 232-241.

50

S. G.SHIRLEY

Ohm, T. (;., Braak, E., Probst, A., and Weindl, A. (1988).Bruin Res. 451, 295-300. Oiiada, N. (1988a). Neuroscience 21, 1003-1012. Onada, N. (1988b). Neurciscic.nce 26, 1013-1022. Ophir, D. ( 1987). Arrh. Otolaqngol., Head Neck Surg. 113, 155- 159. Ophir, D., arid Lancet, D. (1988).Armt. Rec. 221, 754-760. Orbach, 1-1. J . , and Cohen. L. B. (1983).J. Neurosci. 3, 2251-2262. Ottoson, D. (19.56). Artu Phyiol. Scund. 35, Suppl. 122, 1-38. Pace, L., and Lancet. D. (1986). Proc. Null. Acud. Scz. U.S.A. 83,4947-4951. Pace, U . , Hanski, E., Salomon, Y., and Lancet, D. (1985). Nature (London) 316, 255-258. Papka, R. E., and Matulionis, D. H. (1983).CeN Tissue Res. 230, 517-525. Parfenova, E. V. ( 1987). T.sitologiyu 29, 1 144- 1149. Parfenova. E. V., and Etinghof, R. N. (1988). Bzochmistq Biokhzrnzyu ( M o m w ) : (Engl. Truiul.) 53, 435-443. Payne, T. L.. Birch, M. C., and Kennedy, C. E. J., eds. (1986). "Mechanisms in Insect Oltaction." Oxford Univ. Press (Ciarendon), London and New York. Pelosi. P., and Pisanelli, A. M. (1981). Chem. Seiues 6, 3-24. Pelosi, P., Baldaccini. N. E., and Pisanelli, A. M. (1982). Biorhem. J. 201, 245-248. Petiderson, P. E., Greer. C., and Shepherd, G. hI. (1986a). In "Handbook of Behavioral Neurobiology (E. M. Blass, ed.), pp. 163-204. Plenum, New York. Penderson, P. E.. Jastreboff, P., Stewart, U'.B., and Shepherd, G. M. (l986b).J. Cornp. Neurol. 250, 93- 108. Persaud, K. C., DeSimone, J. A,, Getchell, M. L., Heck, G. L.. and Getchell, T. V. (1987). Biochim. Sioplcys. Ada 902, 65-79. Persaud, K. C., Heck, G. L., DeSimone, S. K., Getchell. T. V..and DeSimone, J. A. (19884. B r w h b . Biophj.s . 4 r h 944,49-62, Persaud, K. C., Pelosi, P., and Dodd, G. H. (1988b). Chem. Seirces 13, 231-246. Pevsner,J., Trifletti. K. R.,Strittniatter, S. M.,and Snyder, S. H. (198.5).Proc. Nut/. Acud. Sci. U.S.A. 82, 3050-30.54. Pevsner, J . , Sklar, P. B.. and Snt-der. S. H. (1986). Proc. Nutl. h a d . Sci. U.S.A. 83, 49424946. Pevsner, J., Hwang, P. M., Sklar, P. B., \'enable, J. C., and Snyder, S. H. (1988a). Proc. Nutl. Acud. Sri. L'.S.A. 85, 2383-2388. Pevsner, J., Reed, R. R., Feinstein, P. G.. and Snyder, S. H. (1988b). Science 241,336-339. PfeufFer, E., Mollner, S.. Lancer, D., and Pfeuffer, T. (1989).J. Biol. Chem. 264, 1880318808.

Phillippva, T. M.. Novoselov. V. I., Bystrova, M. F., and Alekseev, S. I. (1988). Bioelertronmgnetirs 9, 34 7-3.54. Pisstrnier, D., Thierry,.J. C., Fabre-Nys, C.. Poindron, P., and Keverne, E. B. (1986). Behau. ,Veut-osri. 100, 361-363. Polak, E. H. (1973).J. Theor. Biol. 40, 469-484. Polak, E. 13.. Shirley. S. G., and Dodo, G. H. (1989).Biocheni. J . 262, 475-478. Potapov, A. A. (1987). Neurophyhlogy 19, 8- 14. Price, S. (1984). Cheni. Serues 8, 341-354. Price, S., arid Turpin, J. (1980).ln"O1faction and 'Taste VII" ( H . Van der Starre, ed.), pp. 65-68. IRL Press, London. Price, S., and Willey, '4. (1987). A n n . 1V.Y. Acad Sci. 510, 561-564. Price, S., and Willey, A. (1988).Biochinc. Biopltw. Actu 965, 127-130. Rafhls, J. A., and Getchell, T. V. (1983). Artul. RPC.206, 87-101. Rambotti, hI. G., Sacccardi, C., Spreca, A,, Aisa, M. C., Giambanco, I., and Donato, K. (1989)..]. Historhem. Cylochm. 37, 1825- 1833.

OLFACTION

51

Reed, C. J., and De Matteis, F. (1989). Biochem. J. 261, 793-800. Reed, C. J., Lock, E. A., and De Matteis, F. (1986). Biochem. J. 240, 585-592. Reed, C. J., J.ock, F. A,, and De Matteis, F. (1988). Biochem. J. 253, 569-576. Rehnberg, B. G., and Schreck, C. B. (1986).J. Comp. Physiol. A 159A, 61-67. Reinken, U., and Schmidt, U. (1986). Exp. Bruin Res. 63, 151-157. Reinken, U., and Schmidt, U. (1987). Nutunuissenschften 74, 555-556. Revial, M. F., Sicard, G., Duchamp, A., and Holley, A. (1982). Chem. Senses 7, 175-190. Revial, M. F., Sicard, G., Duchamp, A., and Holley, A. (1983). Chem. Senses 8, 179-194. Risser, J., and Slotnick, B. M. (1985). Chem. Sewes 10, 410. Robinson, C. J., Shirley S. G., and Dodd, G. H. (1989). Biochem. J. 260, 683-687. Rosser, A. E., and Keverne, E. B. (1985). Neuroscience 15, 1141-1 148. Rosser, A. E., Hokfelt, T., and Goldstein, M. (1986).J. Comp. Neurol. 250, 352-363. Rowley, J. C., 111, Moran, D. T., and Jafer, B. W. (1989).Bruin Res. 502, 387-400. Royet, J. P., Sicard, G. Souchier, C., and Jourdan, F. (1987).Bruin Res. 417, 1-1 1 . Rulli, R. D., and Bruch, R. C. (1987). Chem. Senses 12, 692-693. Russell, Y., Evans, P., and Dodd, G. H. (1989).J. Lip& Res. 30, 877-884. Sakai, M., Yoshida, M., Karhsawa, N., Teramura, M., Ueda, H., and Nagatsu, I. (1987). Expm’entiu 43, 298-300. Sakai, M., Kani, K., Karasawa, N., Yoshida, M., and Nagatsu, I. (1988).BruinRes. 458,335338. Saucier, D., and Astic, L. (1986).Bruin Rex Bull. 16, 455-462. Schild, D. (1988). Biophys. J . 54, 1001-1012. Schild, D. (1989). Exp. Bruin Res. 78, 223-232. Schild, D., and Zippel, H. P. (1986).J. Comp. Physiol. A 158A, 563-571. Schmiedel-Jacob, I., Anderson, P. A. V., and Ache, B. W. (1989).J. Neurophysiol. 61, 9941000.

Schneider, H. (1968). Biochim. Biophys. Actu 163, 451-458. Schofield, P. R. (1988). Trends Neurosci. 11, 471. Schwob, J. E., and Gottlieb, D. I. (1986).J. Neurosci. 6, 3393-3404. Schwob, J. E., and Gottlieb, D. I. (1987). Ann. N.Y. Acad. Sci. 510,597-599. Schwob, J. E., and Gottlieb, D. I. (1988).J. Neurosci. 8, 3470-3480. Schwob, J. E., Farber, N. B., and Gottlieb, D. I. (1986).J. Neurosci. 6, 208-217. Scott, J. W. (1986). E x p m k t i a 42, 223-232. Scott, J. W. (1987). Ann. N.Y. Acud. Sci. 510, 44-48. Scott, J. W., and Harrison, T. A. (1987). In “Neurobiology of Taste and Smell” (T. Finger and W. Silver, eds.), pp. 151-178. Wiley, New York. Scott, J. W., McDonald, J. K., and Pemberton, J. L. (1987).J. Comp. Neurol. 260, 378-391. Seeman, P., Roth, S., and Schneider, R. (1971). Biochim. Biophys. Actu 225, 171-184. Seroogy, K. B., Brecha, N., and Gall, C. (1985).J. Comp. Neurol. 239,373-383. Sharp, F. R., Kauer, J. S., and Shepherd, G. M. (1977).J . Neuro$hysiol. 40, 800-813. Shibuya, T., Aihara, Y., and Tonosaki, K. (1977).In “Food Intake and Chemical Senses”(Y. Katsuki, M. Sato, S. F. Takagi, and Y. Oomura, eds.), pp. 23-32. Univ. of Tokyo Press, Tokyo. Shinoda, K., Shiotani, Y., and Osawa, Y. (1989).J. Comp. Neurol. 284, 362-373. Shipley, M. T., Halloran, F. J., and De la Torre, J. (1985). Bruin Res. 329, 294-299. Shirley, S. G. (1984). I n “Mammalian Semiochemistry” (E. S. Albone, ed.), pp. 243-277. Wiley, New York. Shirley, S. G., and Persaud, K. C. (1991). Semin. Neurosci. 2. Shirley, S. G., and Robinson, C. J. (1988). Trends Neurosci. 11, 532-533. Shirley, S. C., Polak, E., and Dodd, G. H. (1983). Eur.J. Biochem. 132, 485-494.

52

S. G. SHIRLEY

Shirley, S. G., Robinson. C.J., Dickinson, K., Aujla, R., and Dodd, G. H. (1986). Biochem. J. 240, 605-607. Shirley, S. G., Polak, E. H., Mather, R. A., and Dodd, G. H. (1987a). Bi0chem.J. 945, 175184. Shirley, S. G., Polak, E. H., Edwards, D. A,, Wood, M. A,, and Dodd, G. H. (1987b). Bzochtni.,/.245, 185- 189. Shirley, S. G., Robinson, C. J., and Dodd, G. H. (1987~).Biochem. J. 245, 613-616. Sicard, G. (1985). Brain Res. 326, 203-212. Sicard, G., and Holley, A. (1984). Brain Res. 294, 283-296. Sicard, G.,Royet. J. P., and Jourdan, F. (1989).Brain Res. 481, 325-334. Silver, CC‘. L. (1 982). J . Coriip. Phystol. A 148A,379-388. Silver, W. L.. Mason. J. R., Adarns. M. A., and Srneraski, (1986). Bruin Res. 376, 221-229. Simmons, P. A., and Getchell, T. V. (1981).J. Neurofhysiol. 45, 529-549. Skeen, L. G, (1977). Bruin Re.$. 124, 145-153. Sklar, P. B., Anholt, R. R. H., and Snyder, S. H. (1986).J. Biol. Chem. 261, 15538-15543. Sklar, P. B., Anholt, R.R. H., and Snyder, S. H. (1987). Ann. N.Y. Acad. Sci. 510,623-626. Slotnick, B. M., Graham, S., Laing, D. G., and Bell, G. A. (1987). Brain Res. 417,343-346. Slotnick, B. M., Panhuber, H., Bell, G. A,, and Laing, D. G. (1989). Brain Res. 500, 161168.

Snyder, S. H., Sklar, P. B., and Pevsner, J. (1988a).J. B i d . C h m . 263, 13971-13974. Snyder, S. H . , Sklar, P. B., and Pevsner, J. (1988b). In “Molecular Neurobiology of the Olfactory System: Molecular, Membranous and Cytological Studies” (F. L. Margolis and T. V. <;etchell, eds.), pp. 3-24. Plenum, New York. Snyder, S. H., Sklar, P. B., Hwang, P. M., and Pevsner, J. (1989). Trend3 Neuroscz. 12,3.5-37. Stallcup, W., Beasley, L. L., and Levine, J. M. (1985).J. Neurosci. 4, 1090-1101. Stevens, J. C., Cain, W.S., and Burke, P. J. (1988). Chem. Senses 13, 643-654. Stewart, W.B. (1985). Brain Res. 347, 200-203. Stewart, W.B., and Penderson, P. E. (1987). Brain Res. 411, 248-258. Stewart, W.B., Kauer, J. S., and Shepherd, G. M. (1979).J. Conlp. Neurol. 185, 715-734. Stott. W.T., and McKenna, M. J. (1985). Fundant. Apfl. Toxicol. 5, 399-404. Sullivan, R. M., and Leon, M. (1986).DN.Bruin. Rps. 27, 278-282. Sullivan, R. M., Wilson, D. A , , and Leon, M. (l989).,/. Neurosci. 9, 3998-4006. Suzuki, N. (1978).J p n . Symp. Tatre Smell (Supporci), pp. 9-12. Suzuki. N. (1987). A m . A’.K A d . Sci. 510, 647-648. Suzuki, N . (1988). I I I “Receptor and Transduction Mechanisms in Taste and Olfaction” (J. C . Brand, ed.), pp. 239-271. Dekker, New York. Suzuki, N. (1989). C,’hem. S e t ~ w s14,318. Szabo, 7:.Blahser, S.. Denizot, J. P., arid Veron-Ravaille, M. (1987). Neurosci. Lett. 81,2452.50. Taniuchi, hl., Schweilzer, J. B., and Johnson, E. M.(1986). Proc. Natl. Acad. Sci. U.S.A. 83, 19.50- 1954. ‘Ibpzzini, .4., Pelosi, P., Pasqualetto, P. L., and Baldaccini, N. E. (1985). Chem. Senses 10, 45-49, ‘Trapido-Kosenthal. H. G., Gleeson, R. A., Carr. W. E. S., Lambert, S. M., and Milstead, M. L. (1987a). Ann. N.Y. Acud. Sci. 510, 669-672. ‘Trapido-Rosenthal. H. G . , Carr, W. E. S., and Gleeson, R. A. (1987b). J . Neurochem. 49, 1174-1 IN’L. ‘Ti-iotskaya, V. T. [ 1988). Meut~ophysiology20, 432-43.5. Trotier, D. (1986).Pfiueger.s Arch. 407, 589-595. Trotier, D. (1991). SPmin. Neurosci. 2.

OLFACTION

53

Trotier, D., and MacLeod, P. (1983). Brain Res. 268, 225-237. Trotier, D., and MacLeod, P. (1986a). Chem. Senses 11, 674. Trotier, D. and MacLeod, P. (1986b). Brain Res. 374, 205-21 1. Trotier, D., and MacLeod, P. (1987). Ann. N.Y. Acad. Sci. 510, 677-679. van Drongelen, W. (1978).J. Physiol. (London) 277, 423-435. van Drongelen, W., Holley, A., and DGving, K. B. (1978).J. Theor. Biol. 71, 39-48. Verhaagen, J., Oestreicher, A. B., Gispen, W. H., and Margolis, F. L. (1989).J. Neurosci. 9, 683-691. Viereck, C., Tucker, R. P., and Matus, A. (1989).J. Neurosci. 9, 3547-3557. Vodyanoy, V. (1988). In ”Receptor and Transduction Mechanisms in Taste and Olfaction” (J. G. Brand, ed.). Dekker, New York. Vodyanoy, V., and Murphy, R. B. (1983). Science 220,717-719. Vodyanoy, V., and Vodyanoy, I. (1987a). Ann. N.Y. Acad. Sci. 510,683-686. Vodyanoy, V., and Vodyanoy, I. (1987b). Neurosci. Lett. 73, 253-259. Voigt, R., and Atema, J. (1987). Ann. N.Y. Acad. Sci. 510, 692-694. Voigt, J. M., Guengerich, F. P., and Baron, J. (1985). Cancer Lett. 27, 241-247. Vollrath, M., Altmannsberger, M., Weber, K., and Osborn, M. (1985). Dzflerentiation (Berlin) 29,243-253. Wallis, I., Ellis, L., Suh, K., and Pfenniger, K. H. (1985). J. Cell B i d . 101, 1990-1998. Wang, R. T., and Halpern, M. (1980). Am. J. Anat. 157, 399-428. Weinstock, R. S., Wright, H. N., Spiegel, S. M., Levine, M. A., and Moses, A. M. (1986). Nature (London) 322, 635-636. Wellis, D. P., Scott, J. W., and Harrison, T. A. (1989).J. Neurophysiol. 61, 1161-1177. Williams, R., and Rush, R. A. (1988). Brain Res. 463, 21-27. Wilson, D. A., and Leon, M. (1986). SOC.Neurosci. Abstr. 12, 123. Wilson, D. A., Sullivan, R. M., and Leon, M. (1987).J. Neurosci. 7, 3154-3162. Winegar, B. D., Rosick, E. R., and Schafer, R. (1988). Comp. Biochem. Physiol. A 91A, 309316. Wirsig, C. R., and Getchell, T. V. (1986). Brain Res. 385, 10-21. Wright, H. N., Weinstock, R. S., Spiegel, A. M., Levine, M. A., and Moses, A. M. (1987). Ann. N.Y. Acad. Sci. 510, 719-722. Wysocki, C. J. (1979). Neurosci. Behav. Rev. 3, 301-341. Yamagishi, M., Hasegawa, S., Takahashi, S., Nakano, Y., and Iwanaga, T. (1989a). Ann. Otol., Rhinol., Laryngol. 98, 384-388. Yamagishi, M., Nakamura, H., Takahashi, S., Nakano, Y., and Iwanaga, T. (198913).Arch. Histol. Cytol. 52, 375-383. Yamamoto, M. (1976). Arch. Histol. Jpn. 38, 359-412. Yoshii, K., and Kurihara, K. (1989). Synapse 3, 234-238. Zheng, L. M., and Jourdan, F. (1988). Neuroscience 26, 367-378. Zheng, L. M., Caldani, M., and Jourdan, F. (1988). Neuroscience 24, 567-578. Zielinski, B. S., Getchell, M. L., and Getchell, T. V. (1989a). Brain Res. 492, 361-365. Zielinski, B. S., Getchell, M. L., Wenokur, R. L., and Getchell, T. V. (1989b). Anat. Rec. 225, 232-245. Zimmer-Faust, R. K., Tyre, J. E., Michel, W. C., and Case, J. P. (1984). B i d . Bull. (Woods Hole, Mars.) 167, 339-353.

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NEUROPHARMACOLOGIC AND BEHAVIORAL ACTIONS OF CLONIDINE: INTERACTl0NS WITH CENTRAL NEUROTRANSMITTERS Jerry J. Buccafusco Department of Pharmacology and Toxicology Deportment of Psychiatry and Health Behavior Medical College of Georgia Augusta, Georgia 30912 and The Department of Veterans Affairs Medical Center Augusta, Georgia 30912

I. Introduction A. Development of a Novel Antihypertensive Drug B. Current Clinical Indications 11. Receptor Specificity A. Peripheral Sites 8. Central Sites 111. Role of Brain Neurotransmitters in the Antihypertensive Response A. Biogenic Amines 8. Opiates C. Acetylcholine D. Other Neurotransmitters IV. Antiwithdrawal Effects A. Opiate Withdrawal B. A Spinal Cord Model for Opiate Withdrawal C. Other Drugs of Abuse V. Other Pharmacological Actions A. Growth Hormone Secretion B. Inhibition of Cholinesterase Inhibitor Toxicity C. Learning and Memory VI. Summary and Conclusions A. The Diversity of Pharmacological Actions B. Clonidine as a Neuromodulator VII. Future Directions References

55 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 33

Copyright Q 1992 by Academic Press, Inc. All rights of reproduction in any form reserved.

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1. Introduction

A. DEVELOPMENT OF A NOVEL ANTIHYPERTENSIVE DRUG T h e impact of the discovery of a novel pharmacological agent on the understanding of a disease process can often be quite dramatic. A case in point is the discovery of the antipsychotic action of chlorpromazine in the early 1950s. In fact, the use of this compound in the treatment of schizophrenia heralded the beginning of modern psychopharmacology. Inquiries into the mechanism of action of this important compound led to the dopamine hypothesis of schizophrenia, a tenet that continues to provide one of the few durable models of brain dysfunction. T h e discovery of chlorpromazine is interesting in another way, that is, in terms of its similarity in certain respects to clonidine. For example, chlorpromazine has amazed researchers over the years regarding its efficacy in a number of psychological and neurological conditions and in the disturbing number of adverse side effects associated with therapy. This pharmacological diversity is a reflection perhaps of the large number of neurotransmitter receptors the drug interacts with. Thus chlorpromazine can inhibit dopamine, a-adrenergic, serotonergic, cholinergic muscarinic, and histaminergic receptors (Baldessarini, 1990). As with chlorpromazine, clonidine began its pharmacological career inauspiciously. The drug was originally synthesized to be employed as a nasal decongestant. Fortunately, the clinical trials initiated to this end resulted in the discovery of the potent hypotensive and sedating actions of the compound (see Kobinger, 1978). T h e unique properties of clonidine unfolded in the mid-1960s and early 1970s and led to an unimaginable wealth of studies. T h e first of these studies, and perhaps the most important up to the present time, concerned the recognition of the role of central adrenergic receptors in cardiovascular regulation and revealed that the brain could be targeted pharmacologiclly to produce an antihypertensive response. In addition to the ground-breaking research that soon followed the discovery of the drug, another feature of similarity with chlorpromazine is clonidine’s pharmacological diversity. As will become evident from the discussion below, clonidine and related drugs have perhaps an unparalleled number of pharmacological and biochemical properties and have found usefulness in a unprecedented number of clinical syndromes. In the case of clonidine, however, this diversity stems not from classical postsynaptic interactions, but most likely through its ability to interact with a number of neurotransmitter sys-

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tems, central and peripheral, through a presynaptic action. For a most excellent earlier review of clonidine and related compounds, the reader is referred to the works of Schmitt (1977) and Kobinger (1978), two individuals who contributed significantly to our current knowledge of the neuropharmacology of central cardiovascular regulation and the mechanism of the antihypertensive action of clonidine. This review will touch upon some of the critical discoveries, but will focus primarily on later accomplishments in the field. Also, the purpose of this review is to summarize the experiments pointing to the mechanism(s) by which clonidine produces its prominent pharmacological actions.

B. CURRENT CLINICAL INDICATIONS

In Dr. Schmitt’s review of 1977, the currently recognized pharmacological properties of clonidine included the peripheral actions, hypertension, vasoconstriction, contraction of the nictitating membrane, and hyperglycemia. These actions could be ascribed simply to stimulation of peripheral a-adrenergic receptors and might have been expected in view of the original plan to design a compound with nasal decongestant properties. I n addition, and more importantly, his review underscored the recognition that clonidine possessed a number of pharmacological properties that were due to an action on the central nervous system. These included hypotension, inhibition of sympathetic tone, activation of vagal tone, bradycardia, sedation, antinociception, hypothermia, inhibition of water intake, inhibition of food intake, and aggressive behavior. Direct evidence for the central site of clonidine’s blood pressure-lowering ability was first provided by Kobinger and colleagues in the mid-1960s and by Schmitt and colleagues in the early 1970s. The details of these experiments have been extensively reviewed (Schmitt, 1977; Kobinger, 1978). In retrospect, it is somewhat surprising that a drug associated with this list of pharmacological properties would become a useful adjunct to the antihypertensive armamentarium. It was perhaps prophetic that in the first page of his 1977 review, Dr. Schmitt writes, “I believe we are as yet in a preliminary and rudimentary stage of the pharmacological knowledge of this compound.” A considerable number of pharmacological properties of clonidine have been reported during the past 20 years. Many of these are listed in Table I (also see Fielding and Lal, 1981). Several of these actions demonstrated in experimental animals occur at doses not usually achieved following clinical use. Some of these will be discussed in more detail below.

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TABLE I PHARMACOLOGICAL PROPERTIES OF CLONIDINE Site of a adrenoceptor stimulation Effect

Central

Hypotensiodinhibition of sympathetic tone Bradycardialenhancement of vagal tone Sensitize baroreceptor reflex activity Suppression of renin secretion Inhibition of renal tubule fluid absorption Inhibition of phrenic nerve activity M ydriasis Decreased intraocular pressure Reduced secretions from parasympathetic glands Sedation Antinociceptioli Decreased food intake H yperglycemia Decreased drinking H ypothermia Growth hormone secretion Enhanced learning/memory Antiinflammatory Anticonvulsant action

X

Peripheral

X X X

X

X

X X X

X X X X

X X X X X

X X

The only clinical indication recommended by the manufacturer for clonidine is in the treatment of hypertensive disease. Nevertheless, the drug is prescribed routinely for a number of other conditions. These are listed in Table 11.

TABLE 11 REPORTED CLINICALLY USEFULACTIONSOF CLONIDINE A X D RELATED DRUGS Hypertensiodcardiovascular disease Blockage of withdrawal syndromes (opiate, benzodiazepine, nicotine, alcohol) Schizophrenia Acute mania Anxietylpanic attacks Childhood hyperactivity Alzheimer's disease Head injury

Anakgeskdanesthesia Treatment of short stature Tardive dyskinesia Akathiska Social phobia Korsakoffs psychosis Glaucoma Diabetic diarrhea

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II. Receptor Specificity

A. PERIPHERAL SITES The ability of clonidine to activate a-adrenergic receptors was appreciated in early studies utilizing spinal-transected animals or in isolated tissues removed from central nervous control. Although it is commonly appreciated that clonidine and related drugs exhibit some selectivity for a,-adrenergic receptors, in earlier studies it was common to view this receptor subtype as occurring exclusively presynaptically. That is, that a,-adrenergic receptors existed primarily on noradrenergic (sympathetic) nerve terminals, where as a,-receptors were located on sympathetic end organs, the latter mediating vasoconstriction and the former mediating inhibition of norepinephrine release. In fact, the earlier literature and some texts referred to clonidine as an adrenergic antagonist, rather than an agonist, because of its known sympatholytic action. With the advent of selective antagonists, it was soon determined that both types of receptors were localized postsynaptically and that these postsynaptic a,-adrenergic receptors also mediated vasoconstriction (Langer and Hicks, 1984). Receptor binding analysis has provided evidence that the a, receptor subtype is localized primarily at the synapse, whereas the 01, receptor subtype is primarily extrajunctional (Langer and Shepperson, 1982). The direct vasoconstrictor action of clonidine is not generally exploited clinically except in instances in which it is desired to produce vasoconstriction in patients with severe autonomic failure and significant postural hypotension (Robertson et al., 1983). The a, receptor subtype also exists on nerve terminals of preganglionic and postganglionic autonomic nerve endings. Clonidine has been reported to produce weak ganglionic blockade and direct inhibition of norepinephrine release from sympathetic fibers, but this is usually achieved by high, supraclinical doses or at low, nonphysiological stimulation frequencies (Schmitt, 1977; Haeusler, 1976a). In contrast, the release of acetylcholine from parasympathetic nerve terminals in several tissues is quite sensitive to inhibition by clonidine and other a,adrenergic agonists, including norepinephrine (Deck et al., 1971; Werner et al., 1972; Drew, 1978). This parasympatholytic action of clonidine is often encountered as side effects associated with therapy. For example, patients often complain of dry mouth, constipation, and visual disturbances.

JERRY J. BUCCAFC‘SCO

B. CENTRAL SITES Although clonidine is able to block the release of norepinephrine from peripheral sympathetic fibers under certain conditions of stimulation, the ability of the drug to reduce circulating levels of catecholamines and their metabolites is due primarily to its central action (Haeusler, 1976b). As with its peripheral action, clonidine blocks the release of norepinephrine from central catecholaminergic nerve endings. Also, consistent with its peripheral effects, the earlier literature often referred to clonidine as a central noradrenergic antagonist. It is now well accepted that clonidine produces profound effects on brain catecholaminergic fibers, even under resting conditions. Consistent with this action, clonidine inhibits the turnover rate of brain norepinephrine. Again this response is mediated through stimulation of central a,-adrenergic receptors (Rochette et al., 1974, 1982; Reid, 1974; Svensson et al., 1975; Draper et al., 1977; Jhanwar-Uniyal et al., 1985). Whereas these receptors mediating norepinephrine release are probably located on nerve endings, central a,-adrenergic receptors are found on both pre- and postsynaptic aspects of the central adrenergic neuron (Janowsky and Sulser, 1987). Although this inhibition of brain norepinephrine release, in part, underlies the decrease in plasma 3-methoxy-4-hydroxyphenylglycol (MHPG) levels (Charney et ul., 1981; Siever et al., 1984), the decrease in plasma norepinephrine is due mainly to clonidine’s ability to decrease sympathetic outflow through its central actions (Garty et al., 1990). T h e relationship between clonidine’s central inhibition of norepinephrine release and sympathoinhibition is discussed below. It is interesting that despite the diversity of pharmacological actions, most of clonidine’s effects have been ascribed to stimulation of central o r peripheral a,-adrenergic receptors (Table I). Ligand binding techniques have demonstrated a heterogeneous distribution of these receptors in the brain and spinal cord (U’Prichard et al., 1977) and their localization has been employed to help explain some of the pharmacological properties of clonidine. For example, the high density of binding sites in the hypothalamus and medulla may serve as the substrates for clonidine’s sympatholytic and vagomimetic actions. T h e high density of binding in the cortex and locus coeruleus may underlie clonidine’s sedative, antiwithdrawal, and other psychotropic actions. T h e anatomical centers in the brain that mediate all of the pharmacological properties of clonidine have not been elucidated; however, there is agreement as to the site of clonidine’s antihypertensive effect following parenterdl administration. Although microin-jection studies have demonstrated that clonidine can elicit a decrease in blood pressure after administration to various hypo-

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thalamic, medullary, and even spinal sites (Struyker-Boudier et al., 1974; Connor and Finch, 1981; LoPachin and Rudy, 1981; Sinha et al., 1985; Pitts et al., 1986; Elghozi et al., 1989; Buccafusco and Magri’, 1989; also see Isaac, 1980), microinjection of clonidine receptor-blocking agents directly into a restricted region of the rostra1 ventrolateral medulla (RVL) almost completely inhibits the hypotensive action of systemically administered clonidine (Bousquet and Schwartz, 1983; Punnen et al., 1987). T h e RVL contains the origin of the so-called C1 epinephrinecontaining descending pathway. This pathway ultimately makes synaptic contact with the preganglionic sympathetic cell bodies of the intermediolateral spinal cord, and is considered to provide tonic sympathetic activity to these cells (see Reis et al., 1989). Although clonidine is a potent agonist at central and peripheral a,adrenergic receptors, Bousquet and colleagues ( 1984) reported that the hypotensive action of clonidine may be more related to its chemical structure as an imidazole than to its ability to act as an a, agonist. Subsequently, Reis and colleagues (Ernsberger et al., 1987) determined that norepinephrine was not able to completely displace bound [f~-~H]aminoclonidine from a membrane preparation derived from the ventrolateral medulla. This “imidazo1e”-bindingsite was then characterized and demonstrated to exhibit affinity for clonidine equal to the classical a,-adrenergic receptors in the RVL. Imidazole binding was regionally distributed, with frontal cortex exhibiting classical a,-adrenergic binding but virtually no imidazole binding, and the RVL exhibiting a high degree of both imidazole and ap binding. This imidazole-binding site is distinct from subclasses of histamine receptors, but the imidazole structure of histaminergic ligands may explain earlier contentions that clonidine lowered blood pressure, at least in part, through stimulation of histamine (possibly H,) receptors (Karppanen et al., 1977; FriskHolmberg, 1980). The existence of a new putative receptor almost necessitated the search for an endogenous ligand. To this end Atlas and Burstein (1984) reported the partial isolation from calf brain of a clonidine-displacing substance (CDS). The material, which was not one of the known catecholamines, displaced specifically bound clonidine from rat brain membranes. Almost simultaneously then, Bousquet and Reis reported that CDS microinjected into the RVL produced an alteration in arterial pressure. Unfortunately the responses were in opposite directions. Bousquet and colleagues ( 1986) reported that CDS microinjected into the lateral reticular nucleus of the anesthetized cat regularly produced a hypertensive response. In fact, preinjection of CDS inhibited the subsequent fall in blood pressure produced by clonidine injection. They therefore con-

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sidered CDS to be an endogenous clonidine antagonist. Reis and colleagues (Meely et al., 1986) reported that CDS was quite potent in inhibiting the binding of [p-3H]aminoclonidine to receptors derived from bovine RVL. In contrast to Bousquet’s findings, microinjection of CDS into the C1 region of the RVL resulted in a dose-dependent decrease in blood pressure. Later this partially purified CDS was demonstrated to bind with greatest affinity to the imidazole receptor, even more so than clonidine (Ernsberger et al., 1988). T h e opposite effects on blood pressure produced by CDS in the two laboratories is problematic, but not completely unexpected in view of the differences in species employed and slight differences in the purification techniques. Furthermore, Bosquet reported no change in heart rate, whereas Reis reported a significant bradycardia. Clearly, the cardiovascular action of Reis’s CDS is more clonidine-like; however, definite conclusions concerning this issue will await complete isolation and identification of the endogenous molecule. Subsequent experiments with a polyclonal antibody to clonidine have suggested that CDS may contain an aminoimidazoline o r guanidine moiety. I t might be pointed out that the importance of the discovery of this new putative receptor and neurotransmitter could extend beyond the CNS because imidazole receptors sensitive to CDS have been characterized in the renal proximal tubule and CDS has been reported to produce contraction of the rat gastric fundus (Felsen Pt al., 1987; Coupry et al., 1989). It will be interesting to determine whether CDS mimics clonidine’s other actions on second-messenger systems o r in inhibiting the release of other neurotransmitters. In addition to the imidazole receptor, evidence also exists for at least two subclasses of a,-adrenergic receptors, aZAand a2Badrenoceptors. Supporting physiological evidence has included the ability of certain antagonists to discriminate between pre- and postsynaptic a,-adrenergic receptors in different peripheral tissues (Bylund, 1978; Ruff010 et al., 1987; Hieble Pt al., 1988). Regarding central a receptors, there is also evidence to suggest a special variant of the receptor in the spinal cord. Stimulation of spinal CY receptors with clonidine does lead to a decrease in blood pressure; however, blockade of this response by selective antagonists has revealed differences in specificity with respect to peripheral a, receptors (Connor and Finch, 1981). Consistent with this concept is the recent finding that intrathecal injection of clonidine or 6-fluoronorepinephrine in unanesthetized rats was less effective than norepinephrine in inhibiting the pressor response to intrathecal injection of the cholinesterase inhibitor neostigmine (see discussion of central adrenergic-cholinergk interactions in cardiovascular regulation below). In-

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stead, clonidine was more effective in this regard when it was administered intracisternally to stimulate medullary 01 receptors (Buccafusco and Magri’, 1989).

111. Role of Brain Neurotransmitters in the Antihypertensive Response

A. BIOGENIC AMINES T h e possibility that the brain, spinal cord, and peripheral sympathetic nerves play a role in the development and/or maintenance of hypertensive disease is no longer in question. Even in animal models of the disease, including mineralocorticoid, salt-sensitive, and renal ischemia models, in which elevated blood pressure was considered to result exclusively from peripheral factors, it is now apparent that the brain and efferent sympathetic activity is an important contributor to the hypertensive process (de Jong, 1984). Some questions yet to be answered concern the degree of CNS contribution to the pathophysiology, the neuronal structures that are involved, and whether neurochemical abnormalities within such structures can be associated with the disease process. Although great strides in these directions have been made in recent years, no one neuropathological entity has emerged as the causative factor. In fact, many neurotransmitter systems, neurohormones, peptides, and several brain areas have been suggested as playing a role. Such diversity is not surprising in view of the complex neuronal interactions that occur in the regulation of a function as vital as blood pressure. However, the discovery of clonidine and its central mechanism in lowering blood pressure and its ability to stimulate adrenergic receptors immediately pointed an accusing finger at the central adrenergic system as a causative factor in hypertension. One of the first studies to implicate a role for central noradrenergic neurons in the development of hypertension in the now classical model for experimental hypertension, the spontaneously hypertensive rat (SHR), was that performed by Haeusler and colleagues (1972). Their approach was to administer the adrenergic neurotoxin, 6-hydroxydopamine, through the lateral cerebral ventricle to young, prehypertensive rats. Under these conditions the development of hypertension was delayed for several weeks and pressure never rose to the highest levels observed for untreated animals. This finding was subsequently reproduced in several laboratories and in other rat models of hypertension (Doba and Reis, 1974; Erinoff et al., 1975; Haeusler, 1976a; van den

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Buuse, et al., 1984). However, although depletion of brain catecholamines inhibited the development of spontaneous hypertension, a permanent reduction in blood pressure was usually not observed in the adult SHR with established hypertension after similar treatment (Haeusler, l976b; Kubo and Misu, 1981; Woodside et al., 1984). Therefore, the concept evolved that central adrenergic neurons participated in the initiation o r triggering of hypertensive disease, but were not involved in maintaining high blood pressure when it had already fully developed. Nevertheless, these findings appeared to be in concert with the centrally mediated antihypertensive properties of clonidine. A role for central adrenergic neurons in hypertension and in the mechanism of the antihypertensive action of clonidine seem to be inexorably linked. This model would predict that elevated blood pressure in hypertensive disease depends upon increased sympathetic activity resulting from the enhanced activity of central adrenergic neurons. In fact, exaggerated central noradrenergic activity has been observed to be associated with experimental hypertension, although this issue is far from resolved. Increases, decreases, or unchanging catecholamine levels and turnover, enzymes, and metabolites have been reported over the years. Such discrepancies are related in part to the different animal models employed, different brain regions examined, and different stages of’ hypertension when measurements were made. A causal relationship between central adrenergic alterations and increased blood pressure has not been unequivocally demonstrated (Versteeg et al., 1984). Nevertheless, the model has heuristic value and goes on to predict that if hypertension is related to enhanced brain catecholaminergic function, then (1) clonidine should lead to a decrease in the release of brain catecholamines and (2) clonidine’s antihypertensive response should be dependent upon intact, functioning brain catecholaminergic neurons. It is clear that the majority of clonidine’s actions are mediated through stimulation of central a,-adrenergic receptors, but a relationship between changes in norddrenergic neuronal activity and clonidine’s cardiovascular actions has not been established. In fact, the preponderance of evidence suggests that this is not the case. Thus, several investigators have demonstrated that the centrally mediated cardiovascular effects of clonidine require neither functioning brain catecholaminergic neurons nor intact stores of catecholamines (Haeusler, 1974; Kobinger and Pichler, 1974, 1975, 1976; Finch et al., 1975; Warnke and Hoefke, 1977; Reynoldson et al., 1979). Similar results have been obtained with respect to other pharmacological actions of clonidine (Table 111). Though decreased catecholamine release may occur as the result of the presynaptic actions of clonidine, the results of an elegant

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TABLE I11 PHARMACOLOGICAL ACTIONS OF CLONIDINE THAT AREUNAFFECTED BY DEPLETION OF BRAINCATECHOLEMINES Pharmacological actions Decrease arterial blood pressure Inhibit sympathetic nerve activity Enhance reflex vagal bradycardia Produce behavioral depression Produce antinociception Produce mydriasis Inhibit phrenic nerve activity Produce sedation Protect against cholinesterase inhibitor toxicity

References Finch et al. (1975); Warnke and Hoefke (1977); Reynoldson et al. (1979) Haeusler (1974); Kobinger and Pichler (1976) Kobling and Pichler (1975) Florio et al. (1975) Paalzow and Paalzow (1976) Koss and Christensen (1979) von Tauberger et al. (1978) Spyraki and Fibiger (1982) Buccafusco et al. (1988b)

neurochemical and neurohistological study by Haeusler and his coworkers (Lorez et al., 1983) have suggested that this direct inhibitory effect on noradrenergic neuronal activity does not appear to be causally related to the drug’s cardiovascular actions. The conclusion may be stated in an alternative fashion: clonidine stimulates central a,-adrenergic receptors, which are located postsynaptically with respect to adrenergic nerve endings. If this is the case, the question arises as to which neurotransmitter system possesses the relevant clonidine-binding site. Any candidate neurotransmitter should (at least for the simplest model) meet the following criteria: 1. The neurotransmitter should be one that is capable of enhancing or mediating increased sympathetic activity. 2. T h e transmitter system might also be involved in the initiation andlor maintenance of experimental hypertension. 3. Inhibition of the transmitter or blockade of its receptors should lead to a decrease in arterial blood pressure. These criteria do not appear to hold, in general, for central adrenergic systems. Clonidine inhibits the firing rate of noradrenergic neurons, and exogenous administration of the neurotransmitter norepinephrine and related compounds, for the most part, also has been demonstrated to evoke a sympathoinhibitory response. Furthermore, depletion of brain catecholamines or a, receptor blockade produces little change in blood pressure. Nevertheless, although the case for a role for central adrenergic systems in the antihypertensive action of clonidine appears weak,

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it is by no means to be totally discounted. For example, Head and co-

workers (1983) reported a requirement for intact central noradrenergic and serotonergic neurons in the cardiovascular action of clonidine in the unanesthetized rabbit. Although these investigators employed similar selective neurotoxins as with the above-cited studies, they point to having used longer time intervals between toxin and clonidine administration than had been used in earlier studies. Additionally, the use of an unanesthetized animal model and species specificity are to be considered. It might be pointed out, however, that experiments employing reversible acute depletors of central adrenergic neurons such as reserpine and a-methyltyrosine (see Lorez et al., 1983) provide results more in line with those showing no effect of central catecholamine depletion on clonidine’s cardiovascular actions.

B. OPIATES The possibility that endogenous brain opiates or opioidergic pathways play a role in the cardiovascular changes produced by clonidine was

suggested perhaps with the observation that clonidine itself produces a significant analgesic response. T h e analgesic or antinociceptive response to clonidine has been demonstrated in several animals models using a number of different pain paradigms (Schmitt, 1977; Fielding and Lal, 1981). Although the doses required to produce this type of response have often been greater than those required to evoke a significant antihypertensive response, clonidine has been demonstrated to be more potent than morphine on a molar basis (Paalzow, 1974; Paalzow and Paalzow, 1976; Fielding et al., 1978). This action of clonidine has been exploited therapeutically to enhance the effects of general anesthetics, in postoperative control of pain, and to produce localized analgesia following epidural administration (Eisenach el nl., 1989a,b; Kitahata, 1989; Vercauteren et al., 1990). Despite this apparent close relationship between clonidine and the opiate system, studies performed in animal models have demonstrated that brain or spinal a,-adrenergic receptors mediate the antinociceptive action to clonidine. More controversial, however, is the role endogenous opiate systems play in this property of clonidine (Hynes et al., 1983; Tchakarov et al., 1985; Sherman et al., 1988; Tasker and Melzack, 1989; Mastrianni et al., 1989; Porchet et al., 1990). Apparently endogenous opiate involvement is related to the type of pain test employed, the strain of rat, and the subtype of opiate receptor. When this potential relationship between clonidine and opiate receptors was demonstrated, it was reported to be selective for the p subtype (Mastrianni et al., 1989).

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T h e cardiovascular depressant effects of morphine and other p agonists are well known and the potential role of the opioidergic system in the development or maintenance of hypertension has been explored in recent years. Perhaps most pertinent to the present discussion is the issue of whether brain opiate receptor blockade can alter the cardiovascular changes induced by clonidine. Several studies from different laboratories have confirmed the observation that naloxone administration blocks or reverses the clonidine-induced decrease in blood pressure in experimental animals. This inhibitory action is observed more prominently in hypertensive animals (Bennett et al., 1982; Ramirez-Gonzalez et al., 1983; Farsang et al., 1984a; Mastrianni and Ingenito, 1987). T h e results in human studies have been less consistent (Bramnert and Hokfelt, 1983; Rodgers and Cubeddu, 1983; Fuenmayor and Cubeddu, 1986), although the naioxone sensitivity of the clonidine response may be selective for a specific subtype of hypertensive patient (Farsang et al., 1984b). This naloxone-sensitive subset was characterized as exhibiting higher cardiac output, stroke index, plasma renin activity, and plasma epinephrine activity than nonresponders. Also, responders were in general more affected by the clonidine treatment itself. The concept that has developed, therefore, is that in certain forms of hypertension, the antihypertensive action of clonidine is mediated through release of an endogenous opioid transmitter. I n consideration of the simple criteria presented above for a neurotransmitter system to be a candidate for the target of interaction with clonidine in cardiovascular regulation, the endogenous opiate system appears to fall short in this regard. The candidate transmitter should, according to the model, mediate an increase in sympathetic activity. Isolated reports have indicated that under certain conditions morphine and other IJ. agonists increase sympathetic tone and blood pressure, but the predominant view is that morphine and endogenous @-endorphinproduce cardiovascular inhibitory actions (see Ramirez-Gonzalez et al., 1983; Versteeg et al., 1984).Although Met-enkephalin-like opiate activity may be sympathoexcitatory (Versteeg et al., 1984; Fuenmayor and Cubeddu, 1986), biochemical studies have demonstrated the ability of clonidine to cause release of @-endorphinsfrom the brain, in vitro (Versteeg et al., 1984; Mastrianni and Ingenito, 1987). In order to support a role for endogenous opiates in hypertension it must be considered that @-endorphinnormally produces inhibitory sympathetic influences, and that in established hypertension such influence is reduced. The possibility that central a,-adrenergic stimulation causes @-endorphin release, which in turn reduces blood pressure, coupled with the suggestion that such opiate-mediated sympathoinhibition could be reduced in hypertension, deserves merit. This scenario does not quite fit with the

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model presented above, but it does suggest an alternative hypothesis that has been advanced before. That is, that sympathoexcitation associated with hypertension is related to a lack of inhibitory transmitter rather than an excess of an excitatory transmitter. Whereas such a possibility has not been tested for &endorphin, administration of naloxone to hypertensive animal models and to humans has not been associated with dramatic cardiovascular changes (Bramnert and Hokfelt, 1983; Farsang et al., 1984a; Ramirez-Conzalez et al., 1983; Fuenmayor and Cubeddu, 1986; Mastrianni and Ingenito, 1987; Florentino et al., 1987). It could be argued that naloxone is a nonselective opiate antagonist and could potentially eliminate both positive and negative influences of endogenous brain opiates on sympathetic outflow and the cardiovascular system. Finally, to be considered is the ability of naloxone to elevate arterial pressure during shock. This action of naloxone has been attributed to blockade of the central and peripheral cardiovascular depressant effects of the P-endorphin that is released during the development of shock (Holaday, 1983). It is possible that naloxone-induced reversal of clonidine’s antihypertensive action in hypertensive animals may be related to this mechanism, particularly because (3-endorphin release has been associated with clonidine treatment. To further support this possibility is the observation that naloxone appears to more readily reverse the effects of clonidine (that is, once clonidine has produced a decrease in blood pressure) than it is able to prevent a clonidine-induced decrease (Bramnert and Hokfelt, 1983; Farsang et al., 1984b; Shropshire and Wendt, 1983; Ramirez-Gonzalez et al., 1983; Fuenmayor and Cubeddu, 1986; Mastrianni and Ingenito, 1987; Florentino et al., 1987). Nevertheless, the participation of central opiate systems in clonidine’s cardiovascular responses must be considered a viable possibility and may play a greater role in certain types of hypertensive disease o r in certain circumstances of clonidine utilization.

C. ACETYLCHOLINE Studies from several laboratories over the past 40 years have pointed to the critical role of central cholinergic neurons in cardiovascular reg-

ulation. Drugs that activate central muscarinic receptors or enhance neuronally released acetylcholine can elicit a reproducible and significant increase in arterial blood pressure in animals and man (for review, see Brezenoff and Giuliano, 1982; Buccafusco and Brezenoff, 1986). Perhaps the earliest study to point to an interaction between clonidine and central cholinergic neurons involved in cardiovascular regulation

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was reported by Bently and Li (1968).They demonstrated that clonidine pretreatment in rats prevented the centrally mediated hypertensive response to the cholinesterase inhibitor physostigmine. Subsequently, it was demonstrated that pretreatments that increased central cholinergic activity reduced the hypotensive actions of clonidine (Laubie, 1975). Characterization of the clonidine/physostigmine interaction was then carried out by Buccafusco and colleagues, who confirmed the marked inhibitory action of clonidine on the hypertensive response to physostigmine (Buccafusco and Spector, 1980a).They demonstrated that the ability of clonidine to inhibit the pressor response to cholinergic stimulation was selective for indirect-acting agonists such as physostigmine. The pressor response produced by arecoline, a direct muscarinic receptor agonist, was not sensitive to clonidine pretreatment (Buccafusco and Spector, 1980b). This observation was consistent with the finding that clonidine produced a significant reduction in the biosynthesis of acetylcholine in several regions of rat brain, particularly in regions important for cardiovascular regulation, the hypothalamus and medulla (Buccafusco and Spector, 1980c; Buccafusco, 1982, 1984~). The latter action of clonidine was demonstrated to be mediated through stimulation of central a-adrenergic receptors. Also, both clonidine and the related drug a-methyldopa produced decreases in blood pressure and inhibition of brain acetylcholine biosynthesis in hypertensive animals at respective clinically relevant doses (Buccafusco, 1984a,b). Interestingly, blockade of central a-adrenergic receptors with phentolamine reduced the inhibitory effect of clonidine on acetylcholine biosynthesis, but did not itself alter cholinergic dynamics (Buccafusco and Spector, 1980a). In retrospect, it was probably not too surprising to find this ability of clonidine and related drugs to inhibit central cholinergic function, because this relationship was well known for peripheral cholinergic, parasympathetic systems. Thus, stimulation of a,-adrenergic receptors located on parasympathetic nerve endings leads to inhibition of evoked acetylcholine release (see Schmitt, 1977). The central and peripheral “anticholinergic”properties of clonidine may in fact be responsible for many of the side effects reported to occur with therapy (Table IV). Sedation and dry mouth, perhaps the most common side effects, may reflect, respectively, peripheral and central cholinergic blockade. The possibility that inhibition of central cholinergic activity by clonidine leads to its antihypertensive actions as well must be considered. The ability of central muscarinic cholinergic stimulation in several brain regions, including the posterior hypothalamus (Brezenoff and Wirecki, 1970; Brezenoff, 1972; Buccafusco and Brezenoff, 1978, 1979), rostra] medulla (Willette et al., 1984; Giuliano et al., 1989), and spinal

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TABLE IV SIDEEFFECTSTHAT HAVEBEENASSOCIATED WITH CLONIDINE THERAPY OF HYPERI‘ENSIOH

Common

Occasional

Dry mouth Drowsiness Sedation

Constipation Dizziness Headache Fatigue

Rare Gastrointestinal Metabolic CNS

Urinarv retention

cord (Marshall and Buccafusco, 1987; Magri’ and Buccafusco, 1988, 1989; Buccafusco and Magri’, 1990), to produce quite profound increases in blood pressure is consistent with the first of the criteria presented above for a candidate neurotransmitter system. With regard to the second criterion, considerable evidence has been provided in recent years that central cholinergic neurons play an important role in the development and maintenance of experimental hypertension. For example, the pressor response to physostigmine in hypertensive rats has been reported to be significantly greater than similar responses elicited from normotensive controls (Kubo and Tatsumi, 1979; Buccafusco and Spector, 1 9 8 0 ~McCaughran ; el al., 1980; Makari et al., 1989; Buccafusco et al., 1990a). T h e exaggerated pressor response to physostigmine in hypertensive rats suggests that acetylcholine release is enhanced at the site of action of physostigmine, presumably within the rostra1 ventroiateral medulla (Punnen et al., 1986; Giuliano et al., 1989). Although there have been no studies to date that have directly measured brain acetylcholine synthesis or release in hypertensive and normotensive rats, certain markers for the presence of cholinergic neurons have been found to be altered in hypertensive rats. These include the activities of acetylcholinesterase (Yamori, 1976) and choline acetyltransferase (Helke et d., 1980; Edwards et al., 1983) and the density of muscarinic receptors (Edwards et al., 1983; Hershkowitz et al., 1983). The differences, in general, have been positive for hypertensive as compared with normotensive animals. T h e first neurochemical evidence for enhanced brain cholinergic activity in spontaneously hypertensive rats was provided by ‘Trimarchi and Buccafusco ( 1987). They employed the high-affinity uptake of choline by freshly prepared crude synaptosomal fractions as a relative measure of cholinergic activity (Kuhar and Murrin, 1978; Jope, 1979). ‘They reported a significant age-dependent increase in the V,,,, for high-affinity choline uptake into synaptosomal membranes derived

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from the medulla-pons and hypothalamus (but not the striatum) of the SHR compared with age-matched normotensive Wistar Kyoto (WKY) and compared with normotensive outbred normotensive Wistar rats. Also, a highly significant correlation was found between resting systolic blood pressure and the V,,, for high-affinity choline uptake in both the medulla-pons and hypothalamus. This increase was not a consequence of elevated blood pressure and, in fact, the neurochemical changes occurred prior to the development of significant hypertension. The third component of the model would predict that inhibition of brain cholinergic function should lead to an antihypertensive response. Accordingly, it has been demonstrated that depletion of endogenous stores of brain acetylcholine in the adult SHR results in a marked decrease in blood pressure (Buccafusco and Spector, 1980c; Brezenoff and Caputi, 1980; Giuliano and Brezenoff, 1987). Depletion of brain acetylcholine was accomplished in unanesthetized rats by intracerebroventricular administration of hemicholinium-3 (HC-3), a selective inhibitor of high-affinity choline uptake. Inhibition of choline uptake interferes with the rate-limiting step in acetylcholine synthesis and results in a time-dependent depletion of the transmitter (Gardiner, 1961; Rommelspacher et al., 1974; Finberg et al., 1979). Central administration of HC-3 also prevents the pressor response to physostigmine (Brezenoff and Rusin, 1974; Giuliano et al., 1989). Giuliano and Brezenoff (1987) have extended the findings in the SHR indicated above regarding the ability of central injection of HC-3 to lower resting blood pressure. This antihypertensive response was reported to occur in other rat models of hypertension, including deoxycorticosterone acetate (DOCA) salt hypertension and aortic constriction hypertension. In their study, the antihypertensive effectiveness of HC-3 was not a consequence of the hypertension per se, because it could be reproduced in SHRs whose pressure was lowered to normotensive levels by intravenous infusion of vasodilators. Finally, Buccafusco (1984~)reported that central HC-3 treatment and systemic a-methyldopa both produced antihypertensive responses in the SHR. HC-3 pretreatment greatly enhanced the antihypertensive effectiveness of a-methyldopa, a result that alluded to a common mechanism of action. As with depletion of brain acetylcholine, selective blockade of brain muscarinic receptors results in a marked antihypertensive response in the SHR (Coram and Brezenoff, 1983). More recent studies have demonstrated that the M, muscarinic receptor subtype mediates the cardiovascular consequences of muscarinic stimulation and blockade (Pazos et al., 1986; Xiao and Brezenoff, 1988; Sundaram et al., 1988; Giuliano et al., 1989).

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Although the actual role of brain cholinergic systems in hypertensive disease is far from clear, it does fulfill all three criteria put forth above for a candidate neurotransmitter mediating clonidine’s cardiovascular actions. Interestingly, morphine and other receptor agonists also inhibit the release of acetylcholine from peripheral and central cholinergic neurons. This relationship is discussed in more detail below in considering the antiwithdrawal action of clonidine. However, as indicated above, clonidine does enhance brain P-endorphin release. This clonidineopiate interaction could have relevance here because release of the kopiate agonist could also, in theory, result in inhibition of the function of brain cholinergic pressor neurons. Finally, it should be pointed out that the proposed site for clonidine’s antihypertensive action, the rostral ventrolateral medulla, is also the site that mediates the hypertensive response following systemic injection of physostigmine (Punnen et al., 1986; Giuliano et al., 1989). Moreover, this site is the only brain region demonstrated thus far to mediate a significant decrease in blood pressure in norniotensive rats following acetylcholine depletion or muscarinic blockade (Sundaram and Sapru, 1988; Giuliano et al., 1989). Therefore, the role of brain cholinergic pressor systems in normal cardiovascular regulation, in hypertensive disease, in maintaining tonic sympathetic neuronal activity, and in mediating the antihypertensive action to clonidine and related drugs should continue to be considered a viable possibility.

D. OTHERNEUROTRANSMITTERS .4lmost every putative neurotransmitter or modulator discovered in the mammalian central nervous system, when centrally administered, has been demonstrated to alter resting hemodynamics. Perhaps it is not too surprising in view of the importance of the regulation of the cardiovascular system to virtually every physiological function. Less is known, however, regarding the role of these substances in mediating the antihypertensive action of clonidine. However, using the rostral ventrolateral medulla as an example, it is at this important regulatory site that receptors for catecholamines, excitatory and inhibitory amino acids, acetylcholine, opiates, and several other neuropeptides are known to be linked to cardiovascular pathways entering or leaving the region (see Reis et ui., 1989). Because this region may also be the primary site of action for systemically administered clonidine, a potential interaction of this imidazoline with each of these neurotransmitter systems is conceivable.

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IV. Antiwithdrawal Effects

A. OPIATEWITHDRAWAL Despite the relative explosion in research in the area of narcotic analgesics and the opioidergic nervous system during the last decade, very little progress has been made with regard to the pharmacological treatment of opiate addiction. Methadone maintenance continues to provide the mainstay treatment in most clinical settings. However, within the best inpatient environment, rates of retention vary from about 50 to 85% and only 12-28% are likely to remain abstinent for 1-3 years after detoxification Uaffe, 1987). In addition, long-term treatment with methadone can be associated with significant side effects, such as constipation, excessive sweating, decreased sexual function, hormonal abnormalities, and sleep disturbances, effects to which tolerance may not develop. Again, despite the significant research effort in this area, attempts to separate the psychic and dependence properties from the analgesic property of opiates have not led to a clinically useful drug. Furthermore, the endogenous opiate peptides are not devoid of the problems of tolerance or dependence. Also, the earlier suggestion that opiate receptor changes might underlie dependence has not been realized (Redmond and Krystal, 1984). However, during the last few years an alternate pharmacological approach to opiate detoxification has been developed and used quite successfully. Clonidine has been found in several clinical studies to relieve primarily the autonomic components of withdrawal, whereas many of the subjective o r behavioral effects, such as craving, may not be fully reduced in magnitude. Although it is recognized that clonidine and related drugs cannot fully substitute for morphine, and it is not clear whether clonidine can alter relapse rates, clonidine treatment has been employed successfully to facilitate the initiation of opiate antagonist (naltrexone) therapy in heroin addicts (Charney et al., 1982). Perhaps even more importantly, clonidine has provided an important research tool with which to investigate the processes of dependence and withdrawal. Noradrenergic neurons are found in greatest concentration within the pontine locus coeruleus, where they coexist with a high density of opiate receptors (Kosterlitz and Hughes, 1975; Bird and Kuhar, 1977). Both morphine and clonidine can suppress the firing rate of locus coeruleus neurons, although the response to each agonist is mediated through respective opiate and a,-adrenergic receptors (Svensson et al., 1975; Cedarbaum and Aghajanian, 1977), and the apparent firing rate

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of the catecholamine neurons in this brain region increases during morphine withdrawal (Aghajanian, 1978). These findings provide the basis for the noradrenergic hypothesis of narcotic withdrawal. In brief, these data have encouraged the hypothesis that noradrenergic hyperactivity originating within the locus coeruleus is responsible for the behavioral and sympathetic responses associated with narcotic withdrawal (Gold et nl., 1978a,b, 1979, 1980). Although this may be true for the behavioral effects of withdrawal, this hypothesis does not appear to accommodate all the facts concerning the pharmacological properties of clonidine. For example, as discussed above, enhanced central noradrenergic activity is usually associated with sympatho-inhibition, not sympatho-excitation (Schmitt and Fenard, 1971; Heise and Kroneberg, 1973; de Jong et al., 1975; Buccafusco and Brezenoff, 1977). Also, the possibility that clonidine acts indirectly to inhibit norepinephrine release to produce its sympatho-inhibitory actions is contradicted by the fact that depletion of norepinephrine or destruction of norepinephrine nerve terminals does not impair the sympatho-inhibitory actions of clonidine (Haeusler, 1974; Kobinger and Pichler, 1974, 1975, 1976; Finch et al., 1975; Warnke and Hoefke, 1977; Reynoldson et a!., 1979; Lorez et nl., 1983; Buccafusco et a/., 198813). These arguments correspond to those presented above for clonidine’s antihypertensive action and so will not be elaborated here. Finally, studies from several laboratories have demonstrated that significant morphine withdrawal responses could be expressed from brain regions other than the locus coeruleus, including the spinal cord (Wikler and Frank, 1948; Martin and Eades, 1964; Martin et al., 1976; Gilbert and Martin, 1976; Davies, 1976; Yaksh et al., 1977; Delander and Takemori, 1983). The fact that enhanced withdrawal-associated behaviors and sympathetic activity (as measured by postwithdrawal blood pressure increase) can be elicited from cardiovascular centers at different levels of the neuraxis (Marshall and Buccafusco, 1985a) suggests that a redundancy in the mechanism of dependencelwithdrawal may exist. Clonidine can interact with a host of potential neurotransmitters and modulators; however, significant evidence exists to support the concept that cholinergic hyperexcitability underlies the sympatho-excitation associated with narcotic withdrawal. In addition to the fact that certain central cholinergic pathways mediate sympatho-excitation, previous neurochemical and pharmacological data demonstrate that opiates inhibit the function of cholinergic neurons and that during withdrawal an enhanced cholinergic activity is expressed (Redmond and Krystal, 1984; Crossland, 1971; Domino and Wilson, 1973; Pinsky et al., 1973; Vasko and Domino, 1978; Casamenti et al., 1980; Crossland and Ahmed, 1984).

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This enhanced cholinergic activity may occur over pathways similar to those involved in central cardiovascular regulation and hypertension (as discussed above). Another feature linking clonidine and morphine with cholinergic neurons is their ability to inhibit the electrically induced release of acetylcholine from parasympathetic nerve endings (for review, see Kosterlitz et al., 1972; Schmitt, 1977).This shared ability is mediated through stimulation of respective presynaptic opiate and a,-adrenergic receptors. Presynaptic p-opiate and a, receptors both function to inhibit acetykholine release from the superior cervical ganglion by reducing calcium influx during preganglionic nerve stimulation (Araujo and Collier, 1987). Also, the withdrawal syndrome evoked after prolonged exposure of the isolated guinea pig ileum to opiates is inhibited by clonidine. This antiwithdrawal effect of clonidine is mediated through stimulation of presynaptic a, receptors. Likewise, the withdrawal syndrome evoked after prolonged exposure to clonidine in this preparation is inhibited by opiate agonists, mediated through opiate receptors (Chahl, 1985; Araujo and Collier, 1987). Thus clonidine’s antiwithdrawal action in humans may be due at least in part, through its common ability with morphine, to inhibit the function of central as well as peripheral cholinergic neurons. In support of this concept are the findings that the sympatho-excitatory response associated with precipitated withdrawal in the rat is inhibited both by clonidine and by hemicholinium-3, a drug that depletes endogenous stores of brain acetylcholine (Buccafusco, 1983; Marshall and Buccafusco, 1987). Therefore, although several brain neurotransmitter systems have been implicated in mediating the symptoms of narcotic withdrawal (for review, see Redmond and Krystal, 1984),evidence appears most compelling for the role of cholinergic neurons in mediating the autonomic symptoms associated with abstinence. If cholinergic neurons do provide the substrate for certain components of the withdrawal syndrome and the antiwithdrawal effects of clonidine, it would follow that anticholinergic agents should be equally effective as antiwithdrawal agents (as indicated above for the experimental drug hemicholinium-3). In fact, the effectiveness of anticholinergic drugs in this regard is controversial (Redmond and Krystal, 1984).Such inconsistencies may be related to the fact that cholinergic neurons do not participate in mediating all withdrawal signs. Furthermore, drugs such as atropine are nonspecific with regard to muscarinic receptor blockade. For example, the sympatho-excitatory effects of central cholinergic stimulation appear to be mediated through M, rather than M, receptors (Pazos et al., 1986; Xiao and Brezenoff, 1988). In concert with this cardiovascular data, Buccafusco ( 1991) has recently demonstrated that the

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selective M , antagonist pirenzepine did not alter the autonomic or behavioral components of naloxone-precipitated withdrawal in morphinedependent rats. In contrast, the MJM, selective antagonist 4-DAMP was effective in almost completely eliminating the cardiovascular (sympathoexcitatory) signs of withdrawal, and was more effective than clonidine under similar conditions in inhibiting the behavioral signs.

B. A SPINAL C O R D MODELFOR OPIATE WITHDRAWAL

'That the spinal cord could provide a useful model system in which to study the neuronal interaction associated with morphine withdrawal was appreciated more than 30 years ago. For example, Martin and colleagues demonstrated that morphine-dependent spinal-transected dogs exhibit many of the autonomic and somatic reflex signs of withdrawal from spinal cord segments distal to the level of transection (Martin and Eades, 1964; Martin et al., 1976). T h e localized intrathecal, subarachnoid administration of morphine and opiate antagonists in dependent rats also indicates that spinal cord neurons participate in the expression of several characteristic behavioral signs and symptoms of the withdrawal response (Yaksh et al., 1977; Delander and Takemori, 1983). In fact, withdrawal-enhanced activity of sympathetic autonomic neurons originating within the spinal cord has led to identification of these neurons as a possible site for the antiwithdrawal effect of clonidine (Franz et al., 1982). High concentrations of opiate peptides and receptors are localized in spinal laminae 1-111 (H8kfelt et al., 1977; Atweh and Kuhar, 1977). This region also contains a high density of small-diameter (C and A6) primary afferent fibers known to transmit thermal and nociceptive information capable of producing reflex autonomic changes (see Martin, 1982; Carew, 1982). Opiate receptors exist on primary afferent fibers (LaMotte et a/.,1976; Jessell et al., 1979), which, when occupied by morphine, results in a decreased excitability of the terminals (Sastry, 1978; Carstens el ut., 1979). These afferent fibers are generally believed to employ substance P as their neurotransmitter and carry sensory pain information and perhaps information related to local autonomic reflex activity (for review, see Neale and Barker, 1983). However, this concept has been challenged (Bossut et al., 1988).Another possible candidate for carrying afferent nociceptive information is glutamate. The dorsal root and dorsal horn are selectively rich in glutamate, and it has been demonstrated that a subset of spinal neurons directly excited by dorsal root fibers has excitatory membrane receptors activated by L-glutamate (Puil, 1983;

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Hunt, 1983; Blumenkopf, 1988). Irrespective of the nature of the excitatory primary afferent transmitter, other studies support the concept that spinal opiate withdrawal may be mediated over afferent cardiovascular reflex pathways (Marshall and Buccafusco, 1987). An established role for spinal cholinergic neurons has been limited primarily to preganglionic and motor neurons and Renshaw cells. Application of muscarinic agents to spinal motor neurons or sympathetic preganglionic neurons does not elicit excitation (Puil, 1983). If spinal motor neurons are cholinoceptive, the cholinergic receptors activated usually produce inhibitory responses. Collaterals from these neurons activate Renshaw cells (classic recurrent inhibition) that are dually innervated by nicotinic as well as muscarinic receptors (Puil, 1983). Nevertheless, stimulation of spinal muscarinic receptors results in a sympathoexcitatory response (Marshall and Buccafusco, 1987; Magri’ and Buccafusco, 1988). This duality of spinal cholinergic function is interesting because, when morphine withdrawal is precipitated directly from the spinal cord, cholinergic neurons play an inhibitory role. On the other hand, if withdrawal involves higher centers, spinal cholinergic neurons play an excitatory role (Marshall and Buccafusco, 1987). The results of these recent studies suggest the presence of a local spinal cholinergic inhibitory circuit as well as an ascending excitatory pathway (Marshall and Buccafusco, 1987; Magri’ and Buccafusco, 1988). The latter is a novel concept and may serve to explain why excitatory spinal muscarinic synapses have been difficult to identify. Direct evidence for the presence of an ascending vasomotor pathway is not yet available. However, transection of the spinal cord was reported not to significantly alter the level of endogenous spinal acetylcholine below the point of the transection, but did reduce levels above the transection, a finding that is consistent with the presence of primarily ascending spinal cholinergic neurons (Potter and Neff, 1984). For the local inhibitory pathway, it is also interesting that high levels of muscarinic receptors exist in the dorsal horn. After dorsal rhizotomy, the level of choline acetyltransferase (the acetylcholine synthetic enzyme) and the density of muscarinic and opiate receptor binding sites were decreased, suggesting that primary afferent fibers may receive a cholinergic as well as opiate innervation (Gillberg and Wiksten, 1986). These results may help to explain the finding that intrathecal injection of muscarinic agonists produces antinociception and a reduction in substance P levels (Smith et al., 1989). T h e use of postwithdrawal arterial blood pressure increase as an index of the sympathetic component of the narcotic abstinence syndrome dates back to the original studies of Himmelsbach in the 1930s (Himmelsbach, 1937, 1939). Perhaps due to the complexities of main-

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taining patent arterial catheters in animals, the models developed since that time depended primarily on measurement of a spectrum of associated behavioral symptoms as well as changes in body temperature and weight loss. Some of these behavioral changes such as “wet dog shakes” and escape or jumping behavior were characteristic of the opiate withdrawal syndrome in rodents. Studies over the last several years have established that, as in clinical subjects, the withdrawal-associated increase in mean arterial pressure (MAP) can provide a reliable and sensitive index of both the intensity of the sympathetic component of abstinence as well as the degree of physical dependence produced as a result of chronic morphine administration in rats (Buccafusco, 1983; Buccafusco et nf., 1984; Buccafusco and Marshall, 1985; Marshall and Buccafusco, 1985a,b,c, 1987). Furthermore, this autonomic symptom of withdrawal has been employed to predict the antiwithdrawal potential of clonidine and the related drug guanfacine (Buccafusco et al., 1984). Because these experiments can be carried out in unanesthetized, freely moving animals, the classical behavioral signs of morphine withdrawal can also be recorded simuitaneously. Therefore, by combining measures of the cardiovascular and behavioral changes associated with withdrawal, a composite picture of withdrawal can be produced in the rat that is similar in many ways to that originally employed by Himmeslbach in patients. ’The pressor response and behavioral changes associated with morphine withdrawal in dependent rats are quite marked following systemic injection of the narcotic antagonist naloxone (there is no effect on blood pressure and no obvious changes in behavior following similar injection of naloxone in nondependent, naive animals). Behavioral and autonomic signs of withdrawal can be elicited following the intraarterial injection (Marshall and Buccafusco, 1985b,c) of naloxone as well as following the injection of the antagonist into localized areas of the CNS (Marshall and Buccafusco, 1985a). In fact, marked behavioral and autonomic withdrawal symptoms can be obtained following intrathecal (i.t.) injection at the level of the spinal sympathetic (thoracic) outflow (Buccafusco and Marshall, 1985; Marshall and Buccafusco, 1985a, 1987). T h e importance of the spinal cord in mediating autonomic symptoms of withdrawal was underscored by the observation that two consecutive withdrawal responses of equal magnitude could be elicited when the first injection of naloxone was made into the lateral cerebral ventricle and the second was into the intrathecal space. Conversely, once the spinal cord was withdrawn (through i.t. injection of naloxone), a second withdrawal response could not be produced by injection of naloxone via any other route (Marshall and Buccafusco, 1985a). Also, the spinal cord in isolation was capable of mediating a withdrawal response, i.e., a marked

CLONIDINEINEUROTRANSMITTER INTERACTIONS

\

-

79

? ....*..*.,............................ ...........-..........-.--*-. 3

#I)-

_ , ~ ~ ~ ~ _ ~ _ ~ ~ ~ o ~ ~ o - o - - - - ~ = ~ - ~ - -

SPINO-MEDULLARY REFLEX DESCENDING FROM HIGHER CENTERS

FIG. 1. Model of spinal and medullary interactions proposed for cholinergic, opioidergic, and adrenergic receptors in the morphine Withdrawal syndrome; ACh, cholinergic neuron; ENK, enkephalinergic interneuron; a,a-adrenergic (presynaptic)receptors; +, excitatory influence; -, inhibitory influence; ?, unknown neurotransmitter.

pressor response was observed in spinal-transected (Cl), morphine-dependent rats (but not in spinal-transected, nondependent animals) (Marshall and Buccafusco, 1985~). The schematic model of putative neuronal interactions illustrated in Fig. 1 is not meant to indicate actual anatomical connections. Nevertheless, pharmacological manipulations of these potential pathways have provided significant insight into the nature of spinal cord pathways involved in autonomic regulation, in the local generation of opiate withdrawal symptoms, and in the antimorphine withdrawal action of clonidine. In spinal-transected rats (control or morphine dependent), local cardiovascular reflex pathways are intact, because a tail or foot pinch reliably elicits an increase in blood pressure (Marshall and Buccafusco, 1987).

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The fact that such a reflex pathway (see the spinal reflex pathway depicted in Fig. 1) is involved in opiate withdrawal is underscored by the observation that dorsal root deafferentation completely abolished the pressor response to naloxone in morphine-dependent, spinal-transected rats (Buccafusco and Marshall, 1985). T h e interactions proposed for the spinal reflex pathway are also based upon the facts that primary afferent fibers d o not directly innervate the cell bodies of the preganglionic neurons (Puil, 1983) and that enkephalin-containing interneurons can produce pre- and postsynaptic inhibitory influences on primary afferent tertninals in the dorsal horn (Sastry, 1978; Carstens et al., 1979). Chronic stimulation of opiate (enkephalin) receptors could lead to inhibition of the reflex pathway, which in turn would result in the activation of compensator): mechanisms. During withdrawal such compensatory mechanisms would be suddenly unopposed, leading to the withdrawal pressor response. ‘The inhibitory cholinergic influence is suggested by the following findings: (1) Intrathecal injection of naloxone to morphine-dependent rats produces a withdrawal-associated increase in blood pressure (Buccafusco, 1983; Marshall and Buccafusco, 1985b). (2) Intrathecal injection of atropine or hemicholinium-3 enhanced the pressor response to i.t. injection of naloxone in morphine-dependent rats (Marshall and Buccafusco, 1987). Because i.t. injections of cholinergic antagonists d o not alter resting levels of blood pressure, the cholinergic inhibitory system proposed must be activated during withdrawal (or during activation of the reflex pathway). It is also possible that this cholinergic influence may be provided by recurrent collaterals from the preganglionic fibers themselves. If this inhibitory influence is mediated through Renshaw cells, both nicotinic and muscarinic receptors could be involved. (3) The a2 agonist clonidine can produce marked inhibition of central cholinergic function and inhibits the sympathetic component of morphine withdrawal (see above); however, in dependent rats, if naloxone is injected by the intrathecal route, local clonidine pretreatment does not affect the pressor response associated with withdrawal (Buccafusco, 1990). (4) Activation of spinal muscarinic receptors results in an antinociceptive response in the rat (Yaksh et al., 1985; Smith et al., 1989). Spinal antinociception was also associated with a selective decrease in the levels of substance P in the dorsal segment of the cord. Because dorsal root afferents have muscarinic receptors, spinal cholinergic (muscarinic) inhibitory pathways may directly inhibit dorsal root afferent terminals, an action that could account both for the antinociceptive (Smith et al., 1989) and withdrawal-potentiating (Marshall and Buccafusco, 1987) actions of local spinal cholinergic neurons. Finally, as depicted in Fig. 1, dorsal afferent fibers involved in cardiovascular re-

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flexes may activate local or ascending sympatho-excitatory pathways, analogous to stimulation of ascending nociceptive pathways. To support a role for primary afferents mediating a sympathetic component of withdrawal is the study by Sharpe and Jaffe (1986), in which neonatal capsaicin treatment was employed to produce permanent degeneration of primary afferent fibers in the adult rat. Capsaicin treatment significantly attenuated naloxone-precipitated autonomic abstinence symptoms such as salivation, lacrimation, and rhinorrhea in rats implanted with morphine pellets several days earlier. In addition to the local reflex circuit, cholinergic neurons participate in sympatho-excitatory responses in the spinal cord. Buccafusco and coworkers were the first to demonstrate that intrathecal injection of cholinergic agonists in freely moving rats produces a marked pressor response (Magri’ and Buccafusco, 1988). This response is greatest when the injection is made at the lower thoracic level (T10-T 13).The pressor response can be elicited following i.t. injection of low microgram doses of carbachol or neostigmine. The effect of the latter drug is mediated through the release of spinal acetylcholine because it is blocked by both atropine and by local depletion of acetylcholine. As with the local spinal circuit, these cholinergic neurons may not participate in mediating tonic sympathetic activity because spinal treatment with muscarinic-blocking drugs or depletion of spinal acetylcholine does not affect resting blood pressure (Magri’ and Buccafusco, 1988). The trajectory of this pathway is unknown but may involve local spinal interaction or a pathway ascending to higher centers, because (1) spinal section at C 1 blocks the pressor response to i.t. injection of both neostigmine (indirect receptor agonist) and carbachol (direct receptor agonist) (Marshall and Buccafusco, 1987); (2)pretreatment with intracisternal injection of hemicholinium-3 to deplete selectively medullary levels of acetylcholine blocks the pressor response to intrathecal injection of carbachol (Magri’ and Buccafusco, 1989) (note i.t. injection of hemicholinium-3has no effect on the pressor response to carbachol) (Magri’ and Buccafusco, 1988); and (3) intracisternal injection of clonidine (which inhibits the function of central cholinergic neurons) (Buccafusco and Brezenoff, 1977; Buccafusco and Spector, 1980c; Buccafusco, 1984a,b,c; Magri’ et al., 1988) blocks the pressor response to intrathecal injection of neostigmine (Magri’ and Buccafusco, 1989). Therefore, the cholinergic excitatory system is cholinoceptive, but may also be cholinergic (see the spino-medullary reflex pathway, Fig. 1). As with certain cholinergic neurons of higher centers, spinal cholinergic excitatory neurons are inhibited by pretreatment with i.t. injection of a-adrenergic agonists. If this cholinergic system participates in the pressor response to systemic injection of naloxone in mor-

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phine-dependent rats, this pathway could be the substrate underlying the antiwithdrawal effects of intrathecal injection of clonidine (Magri’ and Buccafusco, 1989). T h e nature of the interactions of the central components of this pathway is not understood. T h e pressor response to neostigmine is not altered in the decerebrate rat preparation (Takahashi and Buccafusco, 1989);therefore, the rostral portion of the circuit must be completed within the medullary-pontine region. Although it is not possible to rule out any particular cell group in integrating this portion of the pathway, several sites in the medulla have been implicated in directly modifying sympathetic vasoconstrictor tone. In recent years it has been demonstrated that narrowly defined anatomical regions and cell groups play a direct role in this process. Of particular interest is the rostral ventroiateral pressor region of the medulla. As discussed previously with respect to clonidine’s antihypertensive action, this area projects directly to spinal sympathetic preganglionic neurons and has been demonstrated, using anatomic, physiologic, and pharmacological techniques, to provide tonic descending vasomotor activity (for review, see Calaresu and Yardiey, i988). Although the descending pathway is not cholinergic, the medullary neurons are cholinoceptive, and cholinergic stimulation increases blood pressure, whereas cholinergic inhibition lowers pressure (Sundaram and Sapru, 1988). T h e RVL is also the origin of the C 1 adrenergic descending neurons. Other regions of importance in cardiovascular regulation include the caudal ventrolateral medulla, the origin of A1 noradrenergic fibers, stimulation of which produces sympatho-inhibition. Also, regions in the dorsomedial medulla may have a sympatho-excitatory role (Calaresu and Yardley, 1988). T h e nucleus tractus solitarius (IVTS), site of termination of baroreceptor and chemoreceptor afferents and origin of the A2 noradrenergic cell group, is also a potential site for cholinergic influences on sympathetic control. This region also contains a relatively high density of cholinergic markers (Simon et al., 1985). Finally, the bulbo-spinal (descending) component of the spinomedullary reflex pathway is not cholinergic because the pressor response to intracisternal (medullary) injection of neostigmine is not blocked following intrathecal injection of atropine (note: i.t. injection of neostigmine is blocked by i.t. pretreatment with atropine). Briefly stated, a sympatho-excitatory cholinoceptive pathway exists in the spinal cord, which subsequently activates a locally active or ascending cholinergic system that interacts with a second cholinergic pressor system, probably localized within the medulla. Both the spinal and medullary cholinergic components of this pathway can be inhibited by a,-adrenergic agonists such as clonidine.

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Sympatho-excitatory pathways descending from higher centers (Fig. 1) may also play a role in the morphine withdrawal syndrome. Naloxone-induced blood pressure increases were obtained following injection of the antagonist into the lateral cerebral, or fourth ventricle, in dependent rats (Marshall and Buccafusco, 1985a). The possibility exists that naloxone administered via the cerebrospinal fluid could be potentially distributed to all brain areas. However, two consecutive full withdrawal responses could be obtained in the same animal when the first injection of naloxone was made into the lateral ventricle and the second was made either intrathecally or systemically. Once the spinal cord was withdrawn, a second withdrawal response could not be obtained from any other route of naloxone injection (Marshall and Buccafusco, 1985a). Thus, at least two (supraspinal and spinal) distinct sympatho-excitatory pathways associated with opiate withdrawal exist. The descending pathways may interact with the spinal sympatho-excitatory (spino-medullary) cholinergic pathway (Fig. 1). This possibility is based upon the finding that i.t. injection of anticholinergic drugs inhibits the pressor response to naloxone following systemic injection in dependent rats (Marshall and Buccafusco, 1987). As further evidence for this possibility, i.t. pretreatment with clonidine also inhibits the pressor response to systemic administration of naloxone in dependent rats (Buccafusco, 1990). Therefore, withdrawal sympathetic pathways descending from higher centers must interact with spinal cholinergic pressor pathways susceptible to clonidine inhibition. A direct (noncholinergic) pathway may exist, because cholinergic antagonists do not completely eliminate the withdrawal response to systemic injection of naloxone in dependent rats (Marshall and Buccafusco, 1987). This noncholinergic descending withdrawal pathway may occur independently of the spino-medullary reflex pathway proposed, or it may activate the noncholinergic descending limb (Fig. 1) of the circuit. Although the above data are consistent with the proposed interactions illustrated in the model, many other interpretations are possible. Underscored here are the interactions between cholinergic, adrenergic, and perhaps peptidergic or glutaminergic systems; however, it is most likely that other biogenic amines and opioid peptides play a role in the expression of the withdrawal syndrome. It appears that at least some of these interactions take place in higher brain centers. For example, in several preliminary experiments we have observed that pretreatment with cerebroventricular injection of cholinergic antagonists inhibits the pressor response to systemic administration of naloxone in dependent rats (Buccafusco, 1991). As with the spinal cord data, however, if naloxone is administered by cerebroventricular injection to precipitate with-

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drawal (rather than systemically), pretreatment with cerebroventricular injection cholinergic antagonists enhances the pressor response to naloxone. This feature of cholinergic neurons, inhibiting withdrawal evoked locally and facilitating withdrawal evoked systemically, suggests a redundancy in such interactions and supports the use of the spinal cord as a model in examining the autonomic mechanisms of the opiate withdrawal syndrome. I n summary, current findings in this area are generally supportive of the hypothesis that the spinal cord may provide a good model for the autonomic component of the narcotic withdrawal syndrome, that cholinergic pathways within the spinal cord modify the withdrawal response, and that clonidine may evoke its antiwithdrawal actions at the level of the spinal cord through inhibition with the sympatho-excitatory cholinergic pathway.

DRUGSOF ABUSE C. OTHER As indicated in Table 111, clonidine has been examined clinically for the treatment of withdrawal signs associated with abstinence following continued use of other drugs with abuse liability. Much less is known about the mechanism of clonidine’s actions in these situations than is known about the opiate withdrawal syndrome. For example, it is not clear whether clonidine’s antianxiety properties or its sedative/CNS depressive properties (see Fielding and Lal, 1981) could account for the reported benefit of clonidine in alcohol (Wartenburg, 1983), benzodiazepine (Kunchandy and Kulkarni, 1986), and nicotine withdrawal (Glassman and Covey, 1990). Regarding alcohol abuse, however, clonidine has been reported to interfere with the expression of ethanol intoxication in a mouse model. The inhibitory effect was reported to be comparable to that produced by the imidazodiazepine RO15-45 13, which is known to inhibit ethanol’s effects by a central action (Lister et al., 1989). Clonidine has also been found to be more effective than conventional therapy in treating states of acute alcohol withdrawal, including delirium tremens (Cushman, 1987). Here the effect, as with morphine withdrawal, may be due in part to its central sympatholytic action. Clonidine has a clear antiwithdrawal effect in animal models and, moreover, has been demonstrated to reduce voluntary ethanol consumption in rats (Opitz, 1990). T h e effect is most likely mediated by central a2adrenergic receptors because this latter action of clonidine was mimicked by the related drugs guanfacine and tiamenidine. T h e interaction

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with central a,-adrenergic receptors is also consistent with the ability of the selective antagonist yohimbine to block clonidine’s antiwithdrawal action. Thus, in alcohol abuse clonidine may be a therapeutic adjunct useful on several fronts, including alcohol withdrawal and chronic alcoholism. Nicotine as a drug of abuse is not known for the severity of its withdrawal syndrome. However, the drug is a powerful reinforcer in both animals and man. Clonidine has been shown to be effective in reducing craving in smokers abstinent from tobacco for 24 hr, although a significant effect was obtained in women only, regarding smoking cessation. Quit rates, however, have been reported to be higher than placebo for both genders when clonidine therapy was combined with counseling (see Glassman et al., 1988; Glassman and Covey, 1990). In view of the side effects commonly associated with clonidine therapy, it is not clear how this approach will be continued to be viewed in the future. Nevertheless, the ability of clonidine to produce significant beneficial actions in so many different abstinence syndromes produced in the laboratory and observed in the treatment of addicts is unprecedented for a pharmacological agent and most likely is a reflection of the drug’s ability to interact centrally with so many important neuronal systems.

V. Other Pharmacological Actions

A. GROWTH HORMONE SECRETION Not only is the ability of clonidine to elicit secretion of growth hormone (GH) employed currently in the treatment of short stature (Pintor et al., 1987) and as a test of GH secretory reserve (Gil-Ad et al., 1979), but the level of plasma GH produced by clonidine administration is being used routinely in research protocols to assess the status of central a2adrenergic receptors (Siever et al., 1982; Eriksson et al., 1986). In clinical studies this approach has been applied to investigation of the status of central a,-adrenergic receptors in childhood affective disorders, depression, and schizophrenia. As with its antihypertensive response, clonidine’s GH stimulatory action is mediated through postsynaptic a,adrenergic receptors, that is, central stores of brain catecholamines are not required for expression of this action (Conway et al., 1990). That the a,-adrenergic receptors mediating GH secretion are part of a physiologically relevant pathway is indicated by the ability of central a,-adrenergic

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receptor blockade with yohimbine to inhibit the GH response to insulininduced hypoglycemia (Suri et al., 1990). T h e hypothalamus is a rich source of neurotransmitters, modulators, and peptide hormones. Although the site of the a,-adrenergic receptor innervation involved in this response is not known, clonidine does not appear to act entirely through stimulation of growth hormone-releasing hormone (Suri et al., 1990). In a recent study by Conway and co-workers (1990),the GH response to clonidine in rats was significantly inhibited by metergoline and cyproheptadine, serotonin [5-hydroxytryptamine (5HT,) receptor] antagonists. Depletion of brain serotonin levels using the tryptophan hydroxylase inhibitor parachlorophenylalanine (PCPA) also blocked the GH surge to subsequent injection of clonidine. The site of potential interaction between clonidine and serotonin was identified through experiments in which the selective serotonin neurotoxin, 5,7dihydroxytryptamine, was microinjected into selected hypothalamic nuclei. Selective destruction of serotonergic cells in the arcuate nucleus effectively abolished the GH secretion to clonidine. Despite these suggestive experiments, the authors were reluctant to affirm that the relevant a,-adrenergic receptors were located on arcuate serotonergic neurons. Their trepidation was due in part to the counterargument that clonidine-induced activation of a,-adrenergic neurons generally leads to a reduction in serotonergic neuronal activity. Therefore, if serotonin neurons were interjected in the GH pathway, clonidine should inhibit GH release. Nevertheless, the data are strong for an important interaction between central adrenergic and serotonergic interactions in GH secretion. An alternate scenario was presented in an earlier study by Casanueva and co-workers (1984). These investigators reported that in their clinical studies administration of atropine completely blocked the GH secretory peak following either an arginine o r clonidine stimulus. Atropine treatment also blocked the GH peak following physical exercise. Although the authors did not comment upon a possible adrenergic-cholinergic interaction in GH secretion, they did suggest that a cholinergic synapse may be the final common pathway for a variety of GH stimulants. However, as with the serotonin situation, cholinergic agonists also stimulate GH secretion. For the serotonergic or cholinergic scenarios to be correct, clonidine would have to have an excitatory action on the respective systems. Although clonidine does in general inhibit these systems, this possibility cannot be ruled out. Further studies should be quite fruitful in helping to elucidate this interesting property of clonidine and its relationship with these hypothalamic neurotransmitters.

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B. INHIBITION OF CHOLINESTERASE INHIBITOR TOXICITY In developing antidotes to poisoning by cholinesterase inhibitors, several potential target sites at the cholinergic synapse have been studied, including the postsynaptic receptor and acetylcholinesterase itself. Muscarinic receptor-blocking agents such as atropine have been, and continue to be, the primary means for pharmacological intervention in cases of anticholinesterase poisoning. Oxime reactivators may prove useful when the enzyme is inhibited by an organophosphorus agent. One site that has received much less attention is the presynaptic site, the cholinergic nerve terminal. It is reasonable to expect that reducing acetylcholine release would decrease the toxicity of cholinesterase inhibitors. In fact, inhibitory mechanisms are in place to reduce cholinergic neuronal function in situations of postsynaptic overstimulation. These mechanisms include down-regulation or decreased postsynaptic receptor numbers and decreased release of transmitter from the cholinergic nerve terminal. In cases in which poisoning is slow enough, such adaptive changes allow for significant degrees of cholinesterase inhibition without toxicity and even without overt symptoms. In cases of acute, severe poisoning, such adaptive mechanisms are too slow to prevent the development of toxicity. Acceleration of presynaptic down-regulation by pharmacological agents, therefore, may be of use under such circumstances. The examination of this approach to protection has been limited, perhaps due to a paucity of presynaptic cholinergic blocking agents, or from the fear that such agents might prove highly toxic. Complete blockade of acetylcholine release with botulinum toxin underscores this concern. However, an agent that merely accelerates presynaptic down-regulation without completely inhibiting transmitter release might be of value. In support of this possibility, Buccafusco (1982) first demonstrated a marked protection by clonidine against the manifestations of physostigmine toxicity. In the mouse, clonidine’s protective actions were associated with significant inhibition of the increase in brain acetylcholine induced by the reversible cholinesterase inhibitor. That the mechanism of protection was primarily through central cholinergic and peripheral muscarinic pathways was indicated by the lack of protection afforded by clonidine against the toxic effects of the selective, peripherally acting cholinesterase inhibitor neostigmine. More recent studies employing organophosphate cholinesterase inhibitors (soman and echothiophate) substantiated the physostigmine studies (Aronstam et al., 1986; Buccafusco and Aronstam, 1986, 1987). Moreover, the combined use of

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atropine and clonidine in the pretreatment regimen was found to enhance survival following soman administration. During these experiments it was consistently noted that clonidine-pretreated mice that survived LD,, doses of soman had fewer behavioral side effects than mice that did not receive clonidine. This observation was confirmed in a rat model in which the toxic behavioral effects induced by soman administration were quantitated (Buccafusco et al., 1988a). Again clonidine offered protection against the lethal as well as the toxic behavioral effects of soman. This behavioral toxicity included the development of tremors, hind limb extension, convulsions and jerking motions, chewing, and excessive salivation. Soman also decreased the expression of normal ongoing behaviors such as sniffing, rearing, and general locomotor activity. 'The ability of clonidine to inhibit soman-induced convulsive behavior (Buccafusco et al., l988a, 1989) is consistent with anticonvulsive activity in other animal models (see Baran et al., 1989),a feature of its protection that might help to limit the development of more permanent toxic manifestations. T h e protective effects of clonidine and atropine were usually synergistic, even though clonidine antagonized some of the stereotyped behaviors elicited by protective doses of atropine (Molloy et al., 1986). 'Thus, while enhancing the protective actions of atropine, clonidine also may reduce atropine-induced side effects. The mechanism for this latter effect is yet to be identified. The mechanism of the protective actions of clonidine has been investigated and appears to be more complex than simply inhibition of acetylcholine release. That is, whereas clonidine does produce a marked inhibition of acetylcholine synthesis and release at peripheral and central muscarinic synapses, its other actions on the cholinergic system include a reversible inhibition of acetylcholinesterase and a reversible inhibition of muscarinic receptors (Aronstam et al., 1986; Buccafusco and Aronstam, 1986, 1987). This interaction with the enzyme was observed in both in vivo and in vitru preparations, and in both cases the permanent inhibition of enzyme activity produced by soman was reduced by clonidine treatment. This mode of protection of the enzyme may be similar to that produced by reversible carbamate cholinesterase inhibitors, such as pyridostigmine. Reversible inhibition of cholinesterase essentially protects the enzyme from permanent inactivation by irreversible agents such as soman. Clonidine and many of the tested analogs were also found to interact directly with muscarinic receptors, in an atropine-like manner. Therefore, clonidine and several analogs afford protection against soman poisoning by at least three mechanisms: (1) a reduction in the release of acetylcholine in brain and peripheral muscarinic sites, (2) reversible inhibition of cholinesterase, and (3) blockade of central mus-

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carinic receptors. All of these effects were achieved following administration of protective doses of clonidine. Furthermore, the muscarinic receptor down-regulation that occurs in response to elevated transmitter levels following soman administration is prevented in mice protected with clonidine (Aronstam et al., 1987). This may simply be a reflection of clonidine’s ability to limit acetylcholine release and postsynaptic receptor stimulation. It is not clear to what degree each of these three mechanisms contributes to the ability of clonidine to produce protection against the acute lethal actions of soman. However, several centrally acting a-adrenergic agonists of different chemical structures share this ability with clonidine, and its relative potency as a protective agent was related to its affinity for a-adrenergic binding sites labeled with [3H]clonidine (Buccafusco and Aronstam, 1987). Also, the ability of clonidine to inhibit the biosynthesis of brain acetylcholine is mediated through a-adrenergic receptors (Buccafusco, 1982). It is this action of clonidine, therefore, that appears to predominate in its ability to protect against the acutely toxic actions of soman. It is possible that clonidineinduced protection of cholinesterase from irreversible inactivation by soman may provide a more chronic form of protection, that is, protection long after the clonidine is metabolized or excreted. Along these lines, animals pretreated with clonidine that survive the soman challenge for several days appear behaviorally normal as compared with atropine-pretreated animals or saline-pretreated animals that survive an LD,, dose of soman (Buccafusco et al., 1989,1990b).This apparent difference was observed even though protected animals may have received a higher dose of soman. Initially this finding might not seem noteworthy, because protected animals might be expected to have a better prognosis than nonprotected animals. However, soman is an irreversible inhibitor of acetylcholinesterase, and clonidine is a very shortacting drug, particularly in rodents (Jarrot and Spector, 1978). In fact, animals protected to the same extent as clonidine with high doses (25 mg/kg) of atropine did not appear as behaviorally normal as the clonidine-pretreated animals. In rats (Buccafusco et al., 1988a), 0.5 mg/kg of clonidine produced a degree of protection equivalent to 6 mg/kg of atropine against lethal and soman-induced behavioral effects. T h e ability of a single dose of soman to induce behavioral abnormalities several days later has been reported (Haggerty et al., 1986). In fact, the decrease in spontaneous motor activity induced by an LD,, dose of soman in the rat was observed over 21 days. Such chronic toxic behavioral effects have also been observed following exposure to other organophosphate cholinesterase inhibitors in animals, and in humans following accidental intoxication (for review, see Karczmar, 1984). The

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mechanism for this delayed toxicity is not clear, but it has been reported that significant brain pathology can occur as early as 24 hr following soman administration (Churchill et al., 1985; Pazdernik et al., 1985). It has been suggested that the pathology may result from the severe convulsive activity present soon after soman administration (Samson et al., 1985). Atropine pretreatment is only partially effective in reducing soman-induced convulsive activity and hence delayed brain pathology (Pazdernik et al., 1986). Clonidine pretreatment, however, was more effective than atropine in preventing the occurrence of soman-induced convulsive behavior, and survivors in the clonidine group were less behaviorally impaired than the atropine group (Buccafusco et al., 1989). High doses of atropine do not offer a substantial degree of protection against chronic toxicity, thus of the three mechanisms of clonidine protection stated above, direct muscarinic receptor blockade is probably of minor importance. Because posttreatment with clonidine is not as effective as pretreatment (J. J. Buccafusco, unpublished observation), the ability of clonidine to reduce acetylcholine release is an important contribution to its acute protective actions. The ability to protect cholinesterase from irreversible inactivation (Aronstam et al., 1986; Buccafusco and Aronstam, 1986) may be more important for protection against chronic soman toxicity. The ability of clonidine to produce its antihypertensive response as well as several of its other pharmacological actions is not reduced in animals whose central stores of catecholamines are reduced (see above). Thus, although central a,-adrenergic receptors are required for clonidine’s actions, these receptors are not located on central catecholaminergic nerves. That central cholinergic neurons are endowed with inhibitory clonidine-binding sites is suggested by clonidine’s ability to inhibit brain acetylcholine turnover rates (see above) as well as its ability to offer protection against cholinesterase inhibitor intoxication. It is not yet clear, however, whether central catecholaminergic systems are implicated in the toxic and lethal manifestations of soman intoxication or whether the protection afforded by clonidine involves such pathways. In one study, catecholamine depletion with reserpine or a-methyl-ptyrosine enhanced the toxicity of 2-sec-butylphenyl-N-methylcarbamate and malathion, whereas the monoamine oxidase inhibitor pargyline reduced their toxicities (Takahashi et al., 1987). The implication is that endogenous brain catecholamines provided a tonically active protective mechanism against cholinesterase inhibitor toxicity. Conflicting results were obtained in a subsequent study regarding the role of brain adrenergic systems in clonidine-induced protection. Depletion of brain catecholamines, using effective doses of either reserpine o r a-methyl-p-

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tyrosine as the depleting agent, did not alter soman-induced death or behavioral toxicity (Buccafusco et al., 1988b). This finding was consistent with their earlier report that the centrally acting a-adrenergic blocking drug yohimbine had no effect on physostigmine-induced toxicity (Buccafusco, 1982). Therefore, the findings by Buccafusco and co-workers are consistent with the possibility that either that these a receptors (clonidine-binding sites) are not functionally innervated, or, if they are, that there exists no significant noradrenergic inhibitory tone. It is difficult to reconcile the opposite results obtained by the above two studies; however, it is possible that N-methylcarbamate and malathion have a toxicological profile that differs from that of soman and/or different sites of action. For example, acute intoxication by soman is mediated almost entirely through central mechanisms, whereas the former agents are not as lipid soluble and may include peripheral mechanisms in producing acute intoxication. The results of the two studies are in concert, however, with regard to the finding that elevation of brain catecholamines using pargyline results in significant protection against cholinesterase inhibitor toxicity. In fact, in the study by Buccafusco and co-workers (1988b), pargyline and clonidine were additive in their protective actions. The fact that pargyline-induced protection was time dependent suggests that its mechanism is related to brain catecholamine accumulation. T h e concept that elevation of brain catecholamine levels is inhibitory to cholinergic activity is consistent with the opposing nature of cholinergic and adrenergic functions throughout the nervous system. T h e experiment employing the amine-depleting agents indicated that adrenergic inhibitory tone to central cholinergic systems sensitive to soman is minimal (at least with regard to the cholinergic systems involved in respiratory function and certain behavioral activity), but the results from the pargyline experiments indicate that endogenous catecholamines under the proper circumstances can provide protection against cholinergic overstimulation. It has yet to be determined whether cholinergic neurons sensitive to soman receive inhibitory innervation from adrenergic systems, or whether cholinergic neurons are endowed with inhibitory regulatory receptors sensitive to a2agonists. In the latter case, elevation of catecholamine levels in the extracellular fluid following pargyline administration could potentially activate such receptors. Thus, clonidine administration offers significant protection against soman lethality as well as several toxic manifestations of soman administration, including hypertension, excessive salivation, convulsive behavior, locomotor depression, and a wide profile of stereotyped activity. Clonidine’s protective actions are in many ways similar to those of atropine, but in other ways surpass them. Clonidine was more effective than atropine at reduc-

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ing soman-induced convulsive behavior and at reducing the expression of chronic soman toxicity, possibly because the two are casually related. The mechanism by which clonidine offers protection in the rat was indicated by experiments demonstrating clonidine’s ability to inhibit somaninduced brain acetylcholine accumulation and soman-induced muscarinic receptor down-regulation. Finally, clonidine’s protective actions appear to be independent of central adrenergic neurons; however, elevation of brain catecholamines, like clonidine, offered significant protection against soman toxicity. This finding may be interpreted to indicate that endogenous catecholamine systems may modulate cholinergic neurons under certain conditions and may therefore serve as a protective mechanism against cholinergic overstimulation. Utilization of direct receptor agonists such as clonidine to limit the expression of cholinergic activity is one way to induce this effect. However, it is possible that other, as yet unknown, protectants against cholinesterase inhibitor intoxication could be developed.

C. LEARNING AND MEMORY Alzheimer’s disease is one of the greatest clinical challenges of this century. T h e disease itself, responsible for the institutionalization of a large proportion of the more than 1,000,000 individuals in nursing homes in this country, is characterized by a rapid decline in cognitive and memory abilities, usually in the later years. There is currently no conclusive diagnostic test save autopsy, but post mortem studies have indicated ultrastructural pathological changes characterized by neuritic plaques, paired helical filaments, and granulovacuolar bodies. One of the most consistent abnormalities associated with Alzheimer’s disease is that of brain neurochemistry. Several laboratories throughout the world have reported a significant loss of markers specific for brain cholinergic neurons, including the synthetic enzyme choline acetyltransferase and the degradative enzyme acetylcholinesterase (Bowen et al., 1976; Davies, 1979; Chyle et al., 1983; Younkin et al., 1986; for review, see also Perry arid Perry, 1983; Bartus et al., 1985). T h e degeneration of cholinergic neurons is not global, but there appears to be a rather selective loss of cholinergic fibers originating in the diencephalon, the so-called nucleus basalis of Meynert (Whitehouse et al., 1981; Perry and Perry, 1983; Wilcock et al., 1983; Nagai et al., 1983; Younkin et al., 1986). Although the cholinergic hypothesis of Alzheimer’s disease has been challenged (Palmer et al.. 1986, for example) and it is now generally accepted that several other neurotransmitters o r neurohormones may also play a role

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(Gottfries et al., 1986; Kragh-Sorensen et al., 1986), pharmacological treatments that enhance central cholinergic neurotransmission have generally provided the most reproducible beneficial effects. These have included direct-acting muscarinic receptor agonists such as arecoline, and centrally acting cholinesterase inhibitors such as physostigmine (Beller et al., 1985). Acute beneficial effects have been observed for both classes of drugs, although both have serious limitations with regard to long-term therapy. These include steep, inverted-U-type dose-response curves, tolerance to the beneficial effects, severe side effects, and unpredictable patient response to standard dosing regimens. Recent clinical trials show that a long-acting cholinesterase inhibitor, tetrahydroaminoacridine (THA), produces significant clinical improvement in Alzheimer’s patients, an effect that may continue during long-term treatment (Summers et al., 1986). It is interesting that the class of agent that seems to be of greatest benefit in Alzheimer’s disease is an indirect-acting cholinergic agonist. Thus, even though there exists significant degeneration of cholinergic fibers to cortical limbic structures, inhibition of the degradative enzyme is still able to enhance the effectiveness of neurotransmitter released from the remaining, intact cholinergic nerve endings. Another feature of the Alzheimer’s brain is that, for the most part, the density and affinity of muscarinic receptors are intact (Caufield et al., 1982; Whitehouse and Au, 1986), although the results of recent studies in which specific receptor subtypes (M, o r M,) were examined suggest a selective loss of the M, subtype (Whitehouse and Au, 1986; Quirion et al., 1986). It has been suggested that the M, subtype may be located presynaptically, hence, the loss during cholinergic degeneration (Iversen, 1986). The presynaptic muscarinic receptor, however, generally functions as a negative-feedback receptor, which upon activation inhibits the further release of acetylcholine. It is this presynaptic down-regulation that may be responsible for the tolerance that can develop to prolonged treatment with physostigmine (Becker and Giacobini, 1988). This and the often severe side effects to physostigmine and other cholinergic agonists provide a great clinical challenge. I n addition to the well-known alterations in central cholinergic function, Alzheimer’s disease has also been associated, post mortem, with a decline in brain catecholamines related to neuronal degeneration. Two recent studies typify such findings (Palmer et al., 1987a,b), with the latter study revealing an approximately 50% reduction in markers for norepinephrine innervation of the temporal cortex. Although the ability of centrally acting cholinergic agonists to enhance learning and memory performance in animals and humans has been appreciated for many

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years, it is more recently that central catecholaminergic pathways have been implicated in these processes (Mair et al., 1985; Robbins et al., 1985). Such findings have been strengthened by recent reports that stimulation of central a,-adrenergic receptors by clonidine results in enhanced memory performance in rat (Freedman et d., 1979; Sara et d., 1987) and nonhuman primate (Arnsten and Goldman-Rakic, 1985, 1987) models, and in human patients with Korsakoffs psychosis (McEntee and Mair, 1980; Mair and McEntee, 1986). T h e mechanism by which clonidine elicits these changes is not known, although it is clear that in Korsakoffs patients, clonidine is acting through central pathways, and that the beneficial action of clonidine was greatest in patients exhibiting the greatest loss of CSF markers for norepinephrine (Mair and McEntee, 1986). Perhaps the neural substrate for this effect of clonidine is the significant multiple high-affinity a, receptor binding sites that have been measured in human prefrontal cortex (Carlson and Andorn, 1986). Despite these findings in support of the possibility that clonidine and related drugs could provide symptomatic improvement in human dementias, the results reported in the aged nonhuman primate have been inconsistent. T h e aged monkey has been employed as one of the most relevant models for human dementia, particularly Alzheimer's disease. These animals not only exhibit behavioral deficits in standard delayedresponse paradigms, but neurochemical and morphological changes similar to those seen on autopsy of Alzheimer's disease brain are often observed (for review see Bartus and Dean, 1985). In one series of studies (Arnsten and Goldman-Rakic, 1985, 1987),clonidine improved delayedresponse performance in aged monkeys, an effect that was linked to a,adrenergic receptor stimulation within the prefrontal cortex. However, such beneficial actions were not reproduced in another laboratory (Davis et ul., 1988), from which it was reported that clonidine disrupted performance. T h e latter group speculated that these opposite findings might be related to differences in the testing environment o r paradigm. The earlier work (Arnsten and Goldman-Rakic, 1985, 1987) had employed the presentation of three-dimensional stimuli though the use of a Wisconsin General Test Apparatus (WGTA), whereas the latter study (Davis et al., 1988) had employed presentation of two-dimensional, spatially linked stimuli with an automated apparatus. That the effect of clonidine might be task specific was later demonstrated by Arnsten and Goldman-Rakic (1990), who could not obtain the same magnitude of beneficial effect of clonidine in a delayed nonmatch-to-sample task as they had earlier obtained with the delayed-response task. Recent studies in this laboratory using a completely automated sys-

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tem, but one in which environmental distractors were not completely eliminated (other animals in the same testing room), have confirmed the ability of clonidine to produce a modest but significant improvement in performance of a delayed matching-to-sample task in both young and aged monkeys (Jackson and Buccafusco, 1991). In the presence of low, clinically relevant doses of clonidine, an improvement averaging approximately 10% of baseline performance was obtained with the most effective dose determined for each animal (0.5-10.0 pglkg, i.m). This improvement continued for at least 24 hr following the single administration, much longer than would have been predicted from the known half-life of the drug. By 3 days after administration, responding was again at baseline levels. T h e reason for this chronic improvement is not known, but it could be a consequence of enhanced learning rather than just improved recall or attention. Sedative effects were observed at higher doses ( > l o &kg) in young monkeys, but the aged animals were more sensitive in this regard, with the lO-pg/kg dose producing marked sedation and impairment in performance. The sedative action of clonidine itself does not appear to be the cause of the enhanced performance (Arnsten et al., 1988). Again, despite these promising results in nonhuman primates, clinical trials using clonidine and the related drug guanfacine did not improve the neuropsychological rating of intellectual and memory function in Alzheimer’s patients (Schlegel et al., 1989). This disparity in results regarding Korsakoffs patients indicated above may reflect a more consistent or dramatic loss of cortical noradrenergic innervation associated with the latter disease. It is possible that certain subpopulations of dementia patients, including Alzheimer’s dementia, might still benefit from therapy with central a,-adrenergic agonists. Also, it is generally recognized that Alzheimer’s disease involves alterations in several neurotransmitter systems and it may be necessary to address pharmacologically these multiple deficits.

VI. Summary and Conclusions

A. THEDIVERSITY OF PHARMACOLOGICAL ACTIONS T h e considerable number of pharmacological actions reported for clonidine as indicated in Table I and the substantial number of potential and actual clinical uses (Table 11) are probably unprecedented for a single pharmacological entity. Such a diverse pharmacological profile is undoubtedly a reflection of a diverse mechanism of action. Clonidine

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and related drugs have been demonstrated to interact with classical neurotransmitter systems, the catecholamines, indolamines, cholinergic, opioidergic, and amino acid transmitter systems. For each action of clonidine no single mechanism has been clearly identified as mediating the pharmacological effect. Perhaps the only common link is that most, if riot all, of its actions are mediated through stimulation of a,-adrenergic receptors (Table I). However, even this direct mechanism may be complicated. T h e emergence of multiple clonidine-binding sites as well as the discovery of a novel (nonadrenergic) iniidazole receptor allow for an even greater diversity in its mechanism of action. Also to be considered is the strategic location of clonidine-binding sites. The location of such a site on the neuron soma could have a completely different effect in terms of excitability of the cell if the location of the receptors is the nerve terminals. T h e role of clonidine’s presynaptic actions, particularly regarding central catecholamine systems, has been addressed several times. In many of clonidine’s actions such an inhibitory effect on catecholamine release has not appeared to play an important role. Nevertheless, inhibition of catecholamine release cannot be discounted as an important contributor to clonidine’s long-term actions. It is not yet clear whether “postsynaptic” clonidine-binding sites are innervated by catecholamine nerve terminals. For example, although clonidine’s inhibitory action on central cholinergic transmission is inhibited by a,-adrenergic antagonists, blockade of these receptors in the absence of clonidine does not lead to increased acetylcholine release (Buccafusco and Spector, 1980a). Therefore, either the clonidine receptors on cholinergic neurons are not innervated or the putative noradrenergic tone is quiescent under normal circumstances. In either case, it is possible that such receptors, located on cholinergic or on other systems, continually sample and respond to changes in catecholamine levels in the cerebrospinal fluid. The mutually antagonistic action of norepinephrine and acetylcholine in dually innervated autonomic effector organs is enhanced through presynaptic modulation. That is, norepinephrine overflow during periods of intense sympathetic stimulation can reduce the release of acetylcholine from parasympathetic nerve endings. This relationship between adrenergic and cholinergic neurons may be further amplified in the CNS, which is perhaps more of a closed system than the peripheral circulation. This concept of neurotransmission via the brain extracellular fluid has more recently been termed volume transmission (Fuxe and Agnati, 1991). Volume transmission was originally invoked in part to explain the presence of neurotransmitter receptors at extrasynaptic sites, but the concept may also help to explain the lack of effect of ag-

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adrenergic blocking drugs on cholinergic function indicated above. For example, if central cholinergic activity is indeed modulated by extracellular levels of catecholamines, it is possible that after chronic treatment with clonidine, decreased levels of catecholamines in the cerebrospinal fluid could have an impact on cholinergic neurotransmission. Another factor contributing to the diversity of clonidine’s actions is the nature of the signal transduction processes reported to be activated by the drug. Consistent actions on neuronal cAMP and related systems have been difficult to obtain. Clonidine’s effect on brain cAMP have been demonstrated to be dependent upon the brain region examined and other factors related to specific preparations (see Janowsky and Sulser, 1987; Nakamichi et al., 1987). Part of the problem may also reside in the ability of clonidine to increase intracellular pH (see Marx, 1987). T h e change in pH has been related to clonidine’s ability to enhance Na+ / H exchange as demonstrated in platelets following stimulation of a,-adrenergic receptors. The increase in intracellular pH may trigger the decrease in cAMP often observed following a,-adrenergic receptor simulation in these cells. +

B. CLONIDINE AS A NEUROMODULATOR Despite this diversity of action, clonidine has been employed clinically for several years quite successfully. Although newer related drugs have purported to be associated with less severe side effects than clonidine, in fact, the drug is well tolerated by a large proportion of patients. Perhaps this selectivity is related to the drug’s marked potency for central autonomic pathways. This action is important for its antihypertensive and perhaps its antiwithdrawal properties, the current main clinical applications. In this respect, it is interesting that clonidine is a better antihypertensive agent than it is a hypotensive agent. Thus, in the presence of disease the drug’s actions are more apparent. The inhibitory action of clonidine on neurotransmitter systems is generally modulatory, its effectiveness in inhibiting transmitter release frequency dependent. Unlike direct receptor-blocking agents, clonidine’s modulatory ability could allow for a more subtle degree of regulation. Even in high doses, for example, clonidine does not usually alter the steady-state levels of neurotransmitter and does not completely inhibit the release or synthesis process. Finally, part of clonidine’s selectivity may be related to its singular effectiveness and potency in inhibiting central and peripheral cholinergic muscarinic activity. If this possibility has merit, then

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the role of central cholinergic neurons in mediating many of clonidine’s phartnacological properties as well as in the disease process itself deserves further investigation.

VII. Future Directions

There is no doubt that clonidine will continue to be employed as the experimental drug of choice for investigations examining the effect of central a,-adrenergic receptor stimulation. Some of the clinical indications for the drug will disappear, possibly its use as an antipsychotic or antianxiety agent. However, the use of the drug in withdrawal syndromes will continue to be examined. Clonidine’s analgesic actions will continue to be exploited and may actually surpass the use of opiates in the production of localized spinal analgesia and as a supplement to general anesthetics, before, during, and after surgical procedures. T h e advantage of clonidine and related drugs in this regard is the reduced capacity for producing central respiratory depression and an almost nonexistent abuse liability. Although the use of clonidine as a first-line antihypertensive agent has been supplanted by newer agents targeting peripheral nerves or blood vessels, a reexamination of clonidine’s property as a central sympatholytic agent may be in order. It is generally appreciated that lowering of blood pressure in hypertensive disease per se does always result in protection from secondary cardiovascular complications such as coronary artery disease, left ventricular hypertrophy, vascular damage to the eyes, kidney, and brain, and the production of ventricular arrhythmias (Rosenman, 1989). This kind of toxicity has been ascribed to excessive catecholamine excretion subsequent to enhanced sympathetic activity often associated with essential hypertension. In fact, the use of certain classes of peripherally acting antihypertensive agents may actually enhance sympatho-adrenal outflow though activation of cardiovascular reflex activity o r through plasma sodium and fluid loss (1220, 1989). To date there have been no large or multicenter studies regarding the incidence of cardiovascular-related morbidity or lethality following longterm treatment with a central versus peripheral antihypertensive treatment regimen. If such a study confirms the cardiovascular protective action of clonidine and related drugs, a resurgence in the utilization of this class of antihypertensive agent would be expected. It might be point-

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ed out that clonidine has been demonstrated to provide benefit in chronic heart failure and ischemic heart disease (Giles et al., 1985) and that in hypertensive rats, clonidine significantly reduced blood pressure and heart rate fluctuations, reflecting an enhanced baroreceptor control (Grichois et al., 1990). Finally, clonidine and related drugs may continue to be examined for potential antidementia effects. Despite the disappointing initial results in Alzheimer’s patients mentioned above, clonidine may exhibit beneficial actions in certain categories of human dementia. It might also be borne in mind that central cholinergic agonists such as physostigmine have produced limited benefit in Alzheimer’s patients. The possibility that combined treatment with clonidine and physostigmine may produce an added benefit fits with the observation of multiple neurotransmitter alterations in Alzheimer’s disease. Several studies (discussed above) indicate that clonidine can inhibit many of the pharmacological effects produced by cholinesterase inhibitors, but this does not necessarily imply incompatibility for their use in dementia. The reverse may actually be the case. Although clonidine does inhibit central cholinergic function, studies in our laboratory have demonstrated that this action is selective for certain brain regions. For example, the striatum and hippocampus, which exhibit a high degree of cholinergic innervation, are essentially spared from the inhibitory action of clonidine on their cholinergic neurons (Buccafusco and Spector, 1980a; Buccafusco, 1984a). One hypothesis that is under current investigation in this laboratory is that combined treatment with clonidine and physostigmine results in a greater than additive effect in a delayed matching paradigm in young and aged monkeys. In this situation, it is envisioned that clonidine may have some palliative action of its own on memory and/or learning processes, but may also obviate some of the autonomic side effects associated with physostigmine administration. This selective protective action of clonidine may be due to its inability to interfere with cholinergic function in the hippocampus, a crucial site for memory formation. Along these lines, it has been demonstrated that in rats having lesions of the ascending noradrenergic bundle, the memory-enhancing effect of physostigmine was blocked. A combination of physostigmine and clonidine was required to restore the beneficial effects of central cholinergic stimulation in the lesioned animals (Haroutunian et al., 1990). Thus, clonidine may widen the therapeutic window for physostigmine’s beneficial actions by reducing interfering side effects and by contributing to the relief of noradrenergic deficits associated with aging or the disease process.

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Aghajanian, G. K. (1978). N n t z i r ~(Londoit) 276, 186-188. Araujo. D. M., a n d Collier, B. (1987). Eur.J. Pharvinsol. 139, 179-186. Arnsten, A. F. T., a n d Goldman-Rakic, P. S. (1985). Science 230, 1273-1276. Arnsten, A. F. T., and Goldman-Rakic, P. S. (1987).J. Neural Trarrs. 24, Suppl., 317-324. Arnsten, .4. F. T., a n d Goldman-Rakic, P. S. (1990). Neurobiol. A p g 11, 583-590. Arnsten, A. F. T., Cai, J. X., and Goldman-Rakic, P. S. (1988).J. Neurosci. 8, 4287-4298. Aronstam, R. S., Smith, M. D., and Buccafusco, J. J. (1986). Lfe Scz. 39, 2097-2102. Aronstam, R. S., Smith, M. D., and Buccafusco, J. J. (1987). Neuroscz. Lett. 78, 107-112. .ktkas. D., and Burstein, Y . (1’984). Eur. J . P/m?rrmcol. 144, 287-293. Atweh, S. F., and Kuhar, M. J. (1977). Brain Rex. 124, 53-67. Baldessarini, K.J. ( 1990). I n “The Pharmacological Basis of Therapeutics” (L. S. Goodman, ’4.G. Gilman, T. W. Rall, A. Nies, and P. Taylor, eds.), 8th ed., pp. 383-435. Perganion, New York. Baran, H.. Hortnagl. H., and Hornykiewicz, 0. (1989). Brain Kes. 495, 253-260. Bartus. R. T., and Dean, R. L. (1985). In “Normal Aging, Alzheimer’s Disease and Senile Dementia: Aspects o n Etiology Pathogenesis, Diagnosis and Treatment” (C. G. Gottfries. ed.), pp. 23 1-267. Editions d e I’Universite d e Bruxelles, Brussels. Bartus, R. T., Dean, R. L., Pontecorvo, hf. J., and Flicker, C. (1985). Ann. N.Y. A d . Sci. 444,332-358. Becker, R. E., and (kdcobini, E. (1988). Drug Dm. Res. 12, 163-19.5. 87, 147-151. Beller, S. A., Overall, J. E., a n d Swann, A. C. (1985). Psychopharn~acolog~ Bennett, D. A., DeFeo, J. J., Elko, E. E., and Lal, H. (1982). Drug Deu. Res. 2, 175-179. Bently, G. A., and Li, D. M. F. (1968). E t o : J . Phnwmrol. 4, 124-134. Bird. S. J.. and Kuhar, M. J . (1977). Brain Res. 122, 523-533. Blunienkopf, €3. (1988).Appl. Seurophyiol. 51, 89-103. Bossut, D., Frenk, H., and Mayer, D. J. (1988).Brain Res. 455, 247-253. Bousquet, P., and Schwartz, .I. (1983). Bioc/imz. Phurmacol. 32, 1459- 1465. Bousqriet, P., Feldtnan, J., B L h , R., a n d Schwartz, J. (l984).J. Phannacol. Exf,. The?; 230, 232-236. Bousquet, P., Feldman, J., and Atlas, D. (1986). Eur. J . Phannarol. 124, 167-170. Bowen, D. M., Smith, C., White, P., and Davison, A. N. (1976). Brain 99, 459-496. Bramnert, M.,and Hokfelt, B. (1983). “lctu Piiysiosiol. S c a d . 118, 379-383. Brezenoff, H. E. (1972). Neurophnmzucdo~11, 637-644. Brezenoff, H. E., a n d Caputi, A. C. (1980). Life Sei. 26, 1037-1045. Brezenoff, H. E. ,and Giuliano, R. (1982). Annu. R w . Plzannmol. Toxicol. 22, 341-381. Brezenoff, H. E., a n d Rusin, J. (1974). Etcr. J. Phnntuzrol. 29, 262-266. Brezenoff, H. E., and Wirecki, T. A. (1970). L f e Sci. 9, 99-109. Buccafusco, J. J. (1982).J . Phmrtzurol. Exp. Ther. 222, 595-599. Buccafusco, .J. J. (1983). Phalmacol., B i o c i ~ t ~Belzuu. i. 18, 209-215. Buccafusco, .J. J. (1984a). Dwg Dm. Res. 4, 627-633. Buccafisco, J. J. (1984b). HI-air1 Rrs. Bull. 13, 257-262. Buccafusco, J. J. ( 1 9 8 4 ~ )Hyp~rt~nszor~ . (Dallas) 6, 6 14-62 I. Buccafusco, J. J. (1990). Brain RPS.513, 8-14. Buccafusco, J. J. (1991). Life Sci. 48, 749-756. Buccafusco, J. J., and Aronstam, R. S. (1986).J. Plrannacol. Exp. Ther. 239, 43-47. Buc-c-afusco,J. J., and Aronstam, R. S. (1987). Toxicol. Lett. 38, 67-76.

CLQNIDINEINEUROTRANSMITTER INTERACTIONS

101

Buccafusco, J. J., and Brezenoff, H. E. (1977). Neurophamacology 16, 775-780. Buccafusco, J. J., and Brezenoff, H. E. (1978). Clin. Exp. Hypertern. 1, 219-227. Buccafusco, J. J., and Brezenoff, H. E. (1979). Brain Res. 165, 295-310. Buccafusco, J. J., and Brezenoff, H. E. (1986). Prog. Drug Res. 30, 127-150. Buccafusco, J. J., and Magri’, V. (1989).J. Auton. Nerv. Syst. 28, 133-140. Buccafusco, J. J., and Magri’, V. (1990). Brain Rex Bull. 25, 69-74. Buccafusco, J. J., and Marshall, D. C. (1985). Neurosci. Lett. 59, 319-324. Buccafusco, J. J., and Spector, S. (1980a).J. Phumacol. Exp. Ther. 212, 58-63. Buccafusco, J. J., and Spector, S. (1980b). Experientia 36, 671-672. Buccafusco, J. J., and Spector, S. (1980~). J. Cardiouasc. Phamacol. 2, 347-355. Buccafusco, J. J., Marshall, D. C., and Turner, R. M. (1984). Life Sci. 35, 1401-1408. Buccafusco, J. J., Graham, J. H., and Aronstam, R. S. (1988a). Pharmacol., Biochem. Behav. 29,309-313. Buccafusco, J. J., Aronstam, R. S., and Graham, J. H. (198813). Toxicol. Lett. 42, 291-299. Buccafusco, J. J., Graham, J. H., VanLingen, J., and Aronstam, R. S. (1989). Neurotoxzcol. Terutol. 11, 39-44. Buccafusco, J. J., Makari, N. F., and Hays, A. C. (1990a).Jpn.1. Phamacol. 54, 105-112. Buccafusco, J. J., Heithold, D. L., and Chon, S. H. (1990b). Toxicol. Lett. 52, 319-329. Bylund, D. B. (1978). Trendc Phamacol. Sci. 9, 356-361. Calaresu, F. R., and Yardley, C. P. (1988). Annu. Rev. Physiol. 50, 51 1-524. Carew, T.J. (1982). In “Principles of Neural Science” (E. C. Kandeland and J. H. Schwartz, eds.), pp. 284-292. ElsevierlNorth-Holland, New York. Carlson, M. A,, and Andorn, A. C. (1986). Eur.J. Phamnacol. 123, 73-78. Carstens, E., Tullock, I., Zieglgansberger, W., and Zimmerman, M. (1979). Pfuegers Arch. 379, 143-147. Casamenti, F., Pedata, F., and Corradetti, R. (1980). Neuropharmacology 19, 597-605. Casanueva, F. F., Villanueva, L., Cabranes, J. A., Cabezas-Cerrato, J., and Fernandez-Cruz, A. (1984). J. Clin. Endocrinol. Metabl. 59, 526-530. Caufield, M. P., Straughan, D. W., Cross, A. J., Crow, T., and Birdsall, N. J. M. (1982). Lancet 2, 1277. Cedarbaum, J. M., and Aghajanian, G. K. (1977). Eur. J. Phamacol. 44,375-385. Chahl, L. A. (1985). Br. J. Phamacol. 85, 457-462. Charney, D. S., Menkey, D. B., and Heninger, G. R. (1981). Arch. Gen. Psychiatq 38, 11601180. Charney, D. S., Riordan, C. E., Kleber, H. D., Murburg, M., Braverman, P. et al. (1982). In “Psychopharmacology: The Third Generation of Progress” (H. Y. Meltzer, ed.), pp. 1327-1333. Raven Press, New York. Churchill, L. C., Pazdernik, T. L., Jackson, J. L., Nelson, S. R., Samson, F. E., McDonough, J. H., and McLeod, C. G. (1985). Neurotoxicology 6, 81-90. Connor, H. E., and Finch, L. (1981). Eu7.J. Pharmacol. 76, 97-100. Conway, S., Richardson, L., Speciale, S., Moherek, R., Mauceri, H., and Krulich, L. (1990). Endocrinology (Baltimore) 126, 1022-1030. Coram, W. M., and Brezenoff, H. E. (1983). Drug Dev. Res. 3, 503-516. Coupry, I., Atlas, D., Podevin, R.-A., Uzielli, I., and Parini, A. (1989). J. Phamacol. Exp. Ther. 252, 293-299. Coyle, J. T., Price, D. L., and DeLong, M. R. (1983). Science 219, 1184-1190. Crossland, J. (1971). 1n“Advancesin Neuropharmacology,” (0.Vinaer, 2. Votava, and P. B. Bradley, eds.), pp. 497-523. North-Holland Publ., Amsterdam. Crossland, J., and Ahmed, K. Z. (1984). Neurochem. Res. 9, 351-366. Cushman, P. (1987). Adv. Alcohol Subst. Abuse 7, 17-28.

1 0’2

JERRY 1 . BUCCAFUSCO

Davies, J. (1976). Bratn Res. 113, 31 1-326. Davies, P. (1979). Brain Res. 171, 319-327. Davis, R. E.. Callahan. M.J., and Downs, D. A. (1988). Drug Dev. Res. 12, 279-286. Deck, R., Oberdorf, A,, and Kroneberg, G. (i971). Arzneim-Forsch. 21, 1580-1584. d e Jong, W., ed. (1984). “Handbook of Hypertension,” Vol. 4. Elsevier, Amsterdam. de Jong, W.,Nijkamp, F. P., and Bohus, B. (197.5). Arrh. Int. Pharmacoldyn. Ther. 213, 272284. Delander, G. E., and Takemori, A. E. (3983).Eur. J. Pharmacol. 94, 35-42. Doba, N . , and Reis, D. J. (1974). Circ. RPS.34, 293-301. Domino, E. F., and Wilson. A. E. (1973). Xature (London) 243, 285-286. Draper. A. , J . , Grimes, D., and Redfern, P. H. (1977).J. Pharm. Phurmacol. 29, 175-177. Drew, (;. M .(1978). Llr. J . Phaniucrol. 64,293-300. Edwards, E.. M c l h g h r a n , J. A , , Friedman, R., McNally, W., and Schechter, N. (1983). c h . Exp. Hypertens. A5, 1683-1 702. Eisertact1.J. C . , Castro. M. I., Dewan, D. hi., and Rose, J. C. (l989a).Anesthesiology 70,51-56. Eiset1ach.J. C.. Kauck. R. L., Buzzanell, C., and Lysak, S. Z. (1989b). Anestheszokn~y71,6476.52. Elghozi. J.-L., Head, G. A., Wolf, W. A , , Anderson, C. R., and Korner, P. I. (1989).Brain Re.$. 499, 39-52. Eriksson. E.. Dellborg. $1.. Soderpalnl, B., Carlsson, M., and Nilsson, C . (1986). L f e Scz. 39, 2 103-2 109. ErinofT. L., Heller, A , ?and Oparil, S. (1975). Proc. Soc. Exp. Biol. Med. 150, 748-754. Ernsberger, P., Steely. M.P.. Mann, J. J., and Reis, D. J. (1987). Eur.1. P h a m c o l . 134, 1-13. Ernskrger, P., Meely. hi. P., a n d Reis, D. J. (1988).Brain Res. 441, 309-318. Farsang, C.. Kapcxsi, I., Vajda, L., Varga, K.. Malisak, Z., Fekete, M., a n d Kunos, G. (1984a). Circulation 69, 461-467. Farsang, C.. Varga, K., Vajda, L., Kapocsi, J., Balas-Eltes, A,, and Kunos, G. (1984b). N e i lropepizdr..((Edin/nirgh) 4, 293- 302. Felsen. D., Ernsberger, P., Meely, hi. P., and Reis, D. J. (1987). Eur. J . Phannarol. 142,453Fielding, S., and Lal,H. (1981). ,&fed. Reg. Reu. 1, 97-123. Fielding, W., Wilker. J.. lfynes, M., Szewczak, M.,Novick, W. J., and Lal, H. (1978). J. Phurimcol. Exp. T h . 207, 899-905. Finberg, ,J. P. M.,Buccafusco, J. J . , and Spector, S. (1979). Life Sri. 25, 147-1.56. Finch, L., Buckinghan~,R. E., Moore, R. -1.. and Bucher, T. J. (1975).J. P h m . Pharmacol. 27, 181-186. Florentino, A , , Jimenez. I., Naratijo, J . R., del Carmen Urdin, M., and Fuentes, J . A. (1987). Life Sci. 41, 2445-2453. Florio, V., Bianchi, L.. and Longo, V. G. (1975). h‘europ/tarniacdo~14, 707-714. Franz. D. S . , Hare. B. D., and McCloskey, K. L. (1982). SciPnr~215, 1643-1645. Freedman. 1.. S.,. Backmanit, M. Z., and Quartermain, D. (1979). Pharniacol., Biochen~. Bptuztj. 11, 2.59-263. Frisk-Holmberg, M. (1980).Aria Physid. Srnnd. 108, 191-193. Fuenmayor, N., a r i d Cubeddu, L. (1986). EIN.J. Pharmncol. 126, 189-197. Fuxe. K., and Agrtati, L. F. (1991).“Volume Transmission in the Brain.” Raven Press, New York. Gardiner, J. E. (1961). B i o c h m . J . 81, 297-303. (;art!, 11..Deka-Starosta, A., Chang, P., Kopin, I. J., and Goldstein, D. S. (lYYO).J. Pharrrwrol. Exp. ti it^. 254, 1068- 1075. Gil-Ad, I., ‘lbper, G., and Laron, 2. (1979). Lancet 2, 278-279. Gilbert. P. E.. arid Martin, W. R. (1976). J. Pharmacol. Exp. Tlier. 198, 66-82.

CLONIDINE/NEUROTRANSMITTER INTERACTIONS

103

Giles, T. D., Thomas, M. G., Sander, G. E., and Quiroz, A. C. (1985).J. Cardiouusc. Phar~ L L C O ~8, . S51455. Gillberg, P.-G., and Wiksten, B. (1986).Acta Physiol. Scand. 126, 575-582. Giuliano, &., and Brezenoff, H. E. (1987).J. Cardiovusc. Pharmacol. 10, 113-122. Giuliano, R., Ruggiero, D. A., Morrison, S., Ernsberger, P., and Reis, D. J. (1989).J. Neurosci. 9, 923-942. Glassman, A. H., and Covey, L. S. (1990). Drugs 40, 1-5. Glassman, A. H., Stetner, F., Walsh, B. T., Raizman, P., and Fleiss, J. (1988).JAMA,J. Am. Med. Assoc. 259, 2863-2866. Gold, M. S., Redmond, D. E., and Kleber, H. D. (1978a).Lancet 1,929-930. Gold, M. S., Redmond, D. E., and Kleber, H. D. (1978b). Lancet 2,599-602. Gold, M. S., Byck, R., Sweeney, D. R., and Kleber, H. D. (1979). Biomedicine 30, 1-4. Gold, M. S., Pottash, A. C., Sweeney, D. R., and Kleber, H. D. (198O).JAMA,J. Am. Med. ASSOC. 243, 343-346. Gottfries, C.-G., Bartfai, T., Carlsson, A., Eckernas, S.-A., and Svennerholm, L. (1986). Prog. Neuro-Psychopharmacol. Biol. Psychiatty 10, 405-4 13. Grichois, M.-L., Japundzic, N., Head, G. A., and Elghozi, J.-L. (1990).J. Cardiouasc. Pharmacol. 16, 449-454. Haeusler, G. (1974).A‘aunyn-Schmiedeberg’sArch. Pharmacol. 286, 97-1 1 1. Haeusler, G. (1976a).I n “Regulation of Blood Pressure by the Central Nervous System” (G. Onesti, M. Fernandes, and K. E. Kim, eds.), pp. 53-64. Grune & Stratton, New York. Haeusler, G. (1976b). Naunyn-Schmiedeberg%Arch. Pharmucol. 295, 191-202. Haeusler, G., Finch, L., and Thoenen, H. (1972).Experientia 28, 1200-1203. Haggerty, G. C., Kurtz, P. J., and Armstrong, R. D. (1986).Neurobehau. Toxicol. Teratol. 8, 695-702. Haroutunian, V., Kanof, P. D., Tsuboyama, G., and Davis, K. L. (1990).Brain Res. 507, 261-266. Head, G. A., Korner, P. I., Lewis, S. L., and Badoer, E. (1983).J. Cardiouasc. Pharmacol. 5, 945-953. Heise, A., and Kroneberg, G. (1973).Naunyn-Schmiedeberg’sArch. Pharmacol. 279,285-300. Helke, C., Muth, E. A,, and Jacobowitz, D. M. (1980).Brain Res. 183,425-436. Hershkowitz, M., Eliash, S., and Cohen, S. (1983). Eur. J. Pharmacol. 86, 229-236. Hieble, J. P., Sulpizo, A. C., Nichols, A. J., Willette, R. N., and Ruffolo, R. R. (1988).J. Pharmacol. Exp. Ther. 247, 645-652. Himmelsbach, C. K. (1937). Public Health Rep., Suppl. 125, 1-18. Himmelsbach, C. K. (1939).J. Pharmacol. Exp. Ther. 67, 239-249. Hokfelt, T., Ljundahl, A., Terenius, L., Elde, R., and Nilsson, G. (1977).Proc. Natl. Acad. Sci. U.S.A. 74, 3081-3085. Holaday, 1.W. ( 1983). Annu. Rev. Pharmacol. Toxacol. 23, 54 1-594. Hunt, S. P. (1983). In “Chemical Neuroanatomy,” (P. C. Emson, ed.), pp. 53-84. Raven Press, New York. Hynes, M. D., Atlas, D., and Ruffolo, R.R. (1983).Pharmacol., Biochem. Behav. 19,879-882. Isaac, L. (1980).J. Cardiovasc. Phamacol. 2, S5-19. Iversen, L. L. (1986). Trendr Pharmucol. Sci., Suppl., 44-45. Izzo,J. L. (1989).Am. J. Hypertens. 2, 305s-312s. Jackson, W. J., and Buccafusco, J. J. (1991). Pharmacol., Biochem. Behau. 39, 79-84. Jaffe, J. H. (1987). I n “Psychopharmacology: The Third Generation of Progress” (H. Y. Meltzer, ed.), pp. 1605-1616. Raven Press, New York. Janowsky, A., and Sulser, F. (1987). In “Psychopharmacology: The Third Generation of Pogress” (H. Y. Meltzer, ed.), pp. 249-256. Raven Press, New York. Jarrot, B., and Spector, S. (1978).J. Phamacol. Exp. Ther. 207, 195-202.

104

JERRY J. BUCCAFUSCO

Jessell, T., TSUWO, A., Kanazawa, I., and Ostuka, M. (1979).Brain Kes. 168,247-259. Jhanwar-Uniyal, M.. Levin, B. E., a n d Leibowitz, S. F. (1985).Brain Ke.x 337, 109-316. Jope, R. S. (1979).Brai?i Ke.7. Rev. 1,313-344. Karczmar, A. G. (1984). Fusdum. Appl. Toxicol. 4,SI-Sl7. Karppanen, H.. Paakkari, I., and Paakkari, P. (1977).Eur.J. Pharmacol. 42,299-302. Kitahata. L. M.(1989). An~sth.Analg. (Ctmelnnd) 68,191-193. Kobinger. W. (1978).Rev. Phyiol., Biochem. Plmrmacol. 81, 39-100. Kobinger. W., and Pichler, L. ( 1 974).Eur. J. Phannucol. 27, 15 1 - 154. Kobinger, W., and Pichler, L. (1975).Eur. J. Phormacof. 30, 56-62. Kobinger. \V., and Pichler, L. (1976). Eur. J. Pharnulcol. 40,31 1-320. Koss, M. C., and Christensen. H. D. (1979).A‘aun)?z-Schmeideberg~~ Arch. Pharmacol. 307,45-

50. Kosterlitz, H. by., and Hughes, J. (3975).Lify Sri. 17, 91-96. Kostei-lit/., H. W.. Lord. J. A . H., and Watt, A. J. (1Y72).In “Agonist and Antagonist Actions of Narcotic .Analgesic Drugs,” (H. W. Kosterlitz, € 3 . 0.J. Collier, and J. E. Villareal, eds.), pp. 45-61,Sfacmillan, New York. Kragh-Sorensen. P., Olsen, R. B., Lund, S., Riezen, H. V., and Steffensen, K. (1986).f r o g . .l’eui-rt-P~~rlzcipha~i~~col. Riol. Psyhiatry 10, 479-492. Kulx), T., and Misu, 1’. (19XI).Jpn.J. Pharnzaco/. 31,286-288. Kubo, T., and Tatsumi, M. (1979).Nnuri~n-Schntzedebeg’sArrh. Phrirmncol. 306,81-83. Kuhar, M. J., and blurrin, L. C . (1978).J.Neurochem. 30,15-21. Kunchandy. J., and Kulkarni, S. K. (1986). Ps~chophnnnacologyog,90, 198-202. LaMotte, C:., Pert, C. P.,a n d Snyder, S. H. (1976).Bruin RPS.112, 407-412. Langel. S. Z., and Hicks. P. E. (1984).J . Cardiovaqc. Pharmacol. 6 , S547-558. Langer. S. Z.,and Shepperson, N. B. (1982).Trends f h u m c o l . Sci. 3,440-444. Laubie, M.(197.5).I n “Recent Advances in €Iypertension” (P. Milliez and M. Safar, eds.), pp. 49-59. Societe Aliena, Reims. Lister, R. G., Durcan, M.J., Nutt, D. J., and Linnoila, M. (1989). fj/e Sci. 44, Ill-119. Lopachin, R. M..and Rudy, T. A. (1981). Brain Hes. 224,195-198. Lorrz, H. P., Kiss, D., Da Prada, M., and Haeusler, G. (1983).NautzyIz-Schmiedebergj Arch. Phtirtnncol. 323,307-314.

Magri’, V., and Buccafusco, J. J. (1988).J. Aulon. ‘\‘em. Sytt. 25,69-77. ibfagri’, V., arid B L K C ~ ~ LJ.I SJ.C(1989).J. ~, Aulon. ,Yen’. Syst. 28, 133-140. Magri‘. V., Buccafusco, J. J.. and Aronstani, R. S. (1988).Tuxicol. Appl. Pharmnrol. 95,464473. Mair, R. G., anti AIcEntee, W.J. (1986).P.~ychophumtacolog,88, 374-380. Wair, K. (;., McEntee, W. J., and Zatorre, R. J. (1985).Behnv. Brain Res. 15,247-254. Makari, N. F., Trimarchi, G. R., and Buccafusco, J . J . (1989).il’europhnrmacology 28,379-

386. Marshall. L). C.. and Buccafusco, J. J. (1985a). Brain Res. 329,131-142. iMarshall, D. C.,and Buccafusco, J. J. (1985b).Drng Derl. Res. 5,271-280. Marshall, L). C..and Buccafusco, J. J. ( 1 9 8 5 ~ )Expen‘entzu . 41, 5-6. Marshall, D. C . , and Buccafusco, J. J. (1987).J.,\‘eurosci. 7 , 627-628. 5lartin. ,I. H.(1982).I n “Principles of Neural Science,” (E. C. Kandel and J. H . Schwartz, etis.). pp. 157- 169. Elseevier/Norrh-Holland, New York. Martin, W. R., and Eades, C:. G. (1964). J. Pliarmncol. Exp. Ther. 146, 385-394. Martiri. W’.R., Eades, C. G., Thompson, J. A., Huppler, R. E., and Gilbert, P. E. (1976). J . Phctrn~acol.Exp. Ther. 197,517-532. Marx, J. L. (1987).Science 238,616. Mlastrianni, J. A,, and Ingenito, A. J. (1987).J . P/rarntacol. Exf. Ther. 242,378-387. Mastrianni, J. A., Abhtt, F. \:., and Kunos, G. (1989).Braiu Hes 479,283-289.

CLONIDINE/NEUROTRANSMITTER INTERACTIONS

105

McCaughran, J. A., Murphy, D., Schechter, N., and Friedman, R. (1980).J . Cardiovasc. Pharmacol. 5, 1001-1009. McEntee, W. J., and Mair, R. G. (1980). Ann. Neurol. 27, 466-470. Meely, M. P., Ernsberger, P. R., Granata, A. R., and Reis, D. J. (1986). Life Sci. 38, 1 1 191126. Molloy, A. G., Aronstam, R. S., and Buccafusco, J. J. (1986).Pharmacol., Biochem. Behav. 25, 985-988. Nagai, T., McGeer, P. L., Peng, J. H., McGeer, E. G., and Dolman, C. E. (1983).Neurosci. Lett. 36, 196-199. Nakamichi, H., Murakami, M., Mizusawa, S., Kondo, Y., Sasaki, H., Watanabe, K., Takahashi, A., Sudo, M., and Ono, Y. (1987). Folia Pharmacol. Jpn. 89, 331-337. Neale, J. H., and Barker, J. L. (1983).I n “Handbook of the Spinal Cord,” (R. A. Davidoff, ed.), Vol. 1 , pp. 171-202. Dekker, New York. Opitz, K. (1990). Drug Alcohol Depend. 25, 43-48. Paalzow, G., and Paalzow, L. ( 1 976). Naunyn-Schmiedeberg’s Arch. Pharmacol. 292, 1 19- 126. Paalzow, L. (1974).J . Pharm. Pharmacol. 26, 36 1-363. Palmer, A. M., Procter, A. W., Stratmann, G. C., and Bowen, D. M. (1986).Neurosci. Lett. 66, 199-204. Palmer, A. M., Wilcock, G. K., Esiri, M. M., Francis, P. T., and Bowen, D. M. (1987a).Brain Res. 401, 231-238. Palmer, A. M., Francis, P. T., Bowen, D. M., Benton, J. S., Neary, D., Mann, D. M. A., and Snowden, J. S. (1987b). Brain Res. 414, 365-375. Pazdernik, T. L., Cross, R. C., Giesler, M., Nelson, S., Samson, F., and McDonough, J. (1985). Neurotoxicology 6, 61-70. Pazdernik, T. L., Nelson, S. R., Cross, R., Churchill, L., Giesler, M., and Samson, F. E. (1986). Arch. Toxicol. 9, Suppl., 333-336. Pazos, A., Wiederhold, K.-H., and Palacios, J. M. (1986). Eur. J. Pharmacol. 125, 63-70. Perry, E. K., and Perry, R. H. (1983).In “Alzheimer’s Disease: The Standard Reference” (B. Reisberg, ed.), pp. 93-99. Collier/Macmillan, London. Pinsky, C., Frederickson, R. C. A., and Vasque, A. J. (1973). Nature (London) 242, 59-60. Pintor, G., Loche, R., Cella, S., Puggioni, R., Locatelli, V., and Muller, E. E. (1987).Lancet 1, 1226-1230. Pitts, D. K., Beuthin, I:. C., and Commissaris, R. L. (1986).Eur. J. Pharmacol. 124,67-74. Porchet, H. C., Piletta. P., and Dayer, P. (1990). Life Sci. 46, 991-998. Potter, P. E., and Neff, N. H. (1984). Brain Res. 303, 87-90. Puil, E. (1983). In “Handbook of the Spinal Cord,” (R. A. Davidoff, ed.), Vol. 1 , pp. 105169. Dekker, New York. Punnen, S., Willette, R. N., Krieger, A. J., and Sapru, H. N. (1986). Brain Res. 382, 178184. Punnen, S., Urbanski, R., Krieger, A. J., and Sapru, H. N. (1987).Bruin Res. 422,336-346. Quirion, R., Martel, J. C., Robitaille, Y., Etienne, P., Nair, N. P. V., and Gauthier, S. (1986). Can. J. Neurol. Sci. 13, 503-510. Ramirez-Gonzalez, M. D., Tchakarov, L., Garcia, R. M., and Kunos, G. (1983). Circ. Res. 53, 150-157. Redmond, D. E., Jr., and Krystal, J. H. (1984). Annu. Rev. Neurosci. 7 , 443-478. Reid, J. L. (1974).In “Central Actions of Drugs in Blood Pressure Regulation” (D. S. Davies and J. L. Reid, eds.), pp. 194-203. University Park Press, Baltimore, Maryland. Reis, D. J., Ruggiero, D. A., and Morrison, S. F. (1989). Am. J. Hypertens. 2, 3633-374. Reynoldson, J. A., Head, G. A., and Korner, P. I. (1979).Eur. J. Phannacol. 55, 257-262. Robbins, T. W., Everitt, B. J., Cole, B. J., Archer, T., and Mohammed, A. (1985). Physiol. Psychol. 13, 127-150.

106

JERRY J. BL‘CCAFUSCO

Robenson, D., Goldberg, M. R., Hollister, A. S., Wade, D., and Robertson, R. M. (1983). Ant. J . M e d 74, 193-200. Rochette, L., Bralet, A. M., and Bralet, J. (1974). J. Phannacol. 5, 209-220. Rochette, L., Bralet, A. M., and Bralet, J. (1982). Naunyn-Schmzedebergk Arch. Pharmacol. 319, 40-42. Rodgers, J. F., and Cubeddu, L. X. (1983). Clin. Pharmatol. Ther. 34, 68-73. Rommelspacher, H., Goldberg, A. M., and Kuhar, M. J. (1974). Neurophrmmacology 14, 1015- 1023. Rosemian, R. H. (1989). Am. J. Hy$ertens. 2, 313s-338s. Ruff’olo, R. R., Sulpizio, A. C., Nichols, A. J., DeMarinis, R. M., and Hieble, J. P. (1987). .~aunyri-Sclintiedeberg‘sArch. Pharmacol. 336, 4 15-4 18. Samson, F., Pazdernik, T. L., Cross, R. S., Churchill, L., Giesler, M., and Nelson, S. R. ( 1 985). Pror. We.$!.Phnrniacol. Soc. 28, 183- 185. Sara. S. J., Maho, C.. and Ammassari, M. (1987). Soc. A’eurosri. Abstr. 13, 656. Sastry, B. K. (1978). Eur. J. Pharmacol. 50, 269-273. Schlegel, J . , Mohr, M., Williams, J., hlann, U., Gearing, M., and Chase, T. N. (1989). Clin. L V r u r ~ h a i n u m l12, . 124- 128. Schmitt, H. (1957). I n ”Handbook of Experimental Pharmacology” (F. Gross, ed.), pp. 299-396. Springer-Verlag. New l’ork. Schmitt, H., and Fenard, S. (1971). Arch. I n ! . Pharntacodyn. T h r . 190, 229-240. Sharpe, L. G., andJaffe, J. H. (1986). .Veurosci. Lett. 71, 213-218. Sherman, S. E., Looinis, C. W.,Milne, B., and Cervenko, F. W.(1988). Eur. J. Phannacol. 148, 371-380. Shropshire, A. T., and Wendt, R. I-. (1983).J . Phannacol. Exf. Ther. 224, 494-500. Siever, L. J.. Insell, T. R., Jimerson, D. C., Lake, C. R., Uhde, T. W., Alot, J., and Murphy, D. L. (1982). Psychiafv Hes. 6, 171-183. Siever, L. J.. Uhde, T. W., and Murphy, D. C. (1984). In “Neurobiology of Mood Disorders” (B. Post, ed.), pp. 502-518. Williams & Wilkins, New York. Simon, J. R.. Dimicco. S. K.. Dimicco, j. A., and Aprison, M. H. (1985). Bruin Res. 344,405408. Sinha, J. N., Gurtu, S.,Sharma, D. K.. and Bhargava, K. P. (1985). A‘aunyn-Schmzedegerg$ Arch. Pharmnarol. 330, 163-168. Smith, M. D., Ymg, X., Nha, J.-Y., and Buccafusco, J. J. (1989). Li/e Sci. 45, 1255-1261. Spyraki, C., and Fibiger, H. C. (1982).J. A‘eitml Trantni. 54, 153-163. Struyker-Boudier, H. A. J., Smeets, G. W. hl., Brouwer, G. M., and von Rossum, J. M. (1974). Neurophan,racology 13, 837-846. Summers, M! K., hlajovski, L. V., Marsh, G. M., Tachiki, K., and Kling, A. (1986). N. Engl. J . Med. 315, 1241-1245. Sundarani, K.?a n d Sapru, H. (1988). J. Aufon. Nem. Sysf. 22, 221-228. Sundarani, K., Krieger, A. J., and Sapru. H. (1988). Brain Re$. 449, 141-149. Suri, D., Hindmarsh, P. C., Brain, C. E., Pringle, P. J., and Brook, C. G. D. (1990). Clin. Endoct+nol. (Oxford) 33, 399-406. Svensson, T. H., Bunney, B. S., and Aghajanian, G. K. (1975). Brain Rex. 92, 291-306. Takahashi, I-I., and Buccafusco, (1989). Soc. Neurosci. Absfr. 15, 597. ‘Takahashi. H . , T m a k a , J., Tsuda, S., and Shirasu, Y. (1987). Furdam. APpl. Toxicol. 8,415422. T a k e r , R. A. R.. a n d Melzack, R. (1989). Lije Sci. 44, 9-17. Tchakarov, L., Abbort, F. V., Rantirez-Gonzalez, hl. D., and Kunos, G. (1985). Bruin Res. 328, 33-40. Trirnarchi, G. R . , a n d Buccafusco, J. J. (1987). h’eurochem. Res. 12, 247-252.

CLONIDINE/NEUROTRANSMITTER INTERACTIONS

107

U’Prichard, D. C., Greenberg, D. A., and Snyder, S. H. (1977). Mol. Pharmucol. 13,454476. van den Buuse, M., deKloet, E. R., Versteeg, D. H. G., and de Jong, W. (1984). Brain Res. 301, 221-229. Vasko, M. R., and Domino, E. F. (1978).J . Pharmucol. Exp. Ther. 207, 848-858. Vercauteren, M., Lauwers, E., Meert, T., De Hert, S., and Adriaensen, H. (1990).AnaestheS ~ U45, 531-534. Versteeg, D. H. G., Petty, M. A,, Bohus, B., and de Jong, W. (1984). In “Handbook of Hypertension” (W. de Jong, ed), Vol. 4, pp. 398-430. Elsevier, Amsterdam. von Tauberger, G., Thoneick, H.-U., and Dulme, H.-J. (1978). Arzneim.-Forsch. 28, 651654. Warnke, E., and Hoefke, W. (1977). Arnzeim.-Forsch. 27, 2311-2313. Wartenburg, A. A. (1983).JAMA, J . Am. Med. Assoc. 9, 1271. Werner, U., Starke, K., and Schumann, H. J. (1972).Arch. Int. Pharmacodyn. Ther. 195,282290. Whitehouse, P. J., and Au, K. S. (1986). Prog. Neuro-Psychophurmacol. Biol. Psychiatry 10, 665-676. Whitehouse, P. J., Price, D. L., Clark, A. W., Coyle, J. T., and DeLong, M. R. (1981).Ann. Neurol. 10, 122-126. Wikler, A., and Frank, K. (1948).J . Phunnacol. Exf. Ther. 94, 382-400. Wilcock, G. K., Esiri, M. M., Bowen, D. M., and Smith, C. C. T. (1983). AMl. Neurobiol. 9, 175-1 79. Willette, R. N., Punnen, S., Krieger, A.]., and Sapru, H. N. (1984).J . Phurmacol. Exp. Ther. 231, 457-463. Woodside, J. R., Beckman, J. J., Althaus, J. S., and Miller, E. D. (1984). Anesth. Analg. (Cleveland) 63, 482-488. Xiao, Y.-F., and Brezenoff, H. E. (1988). Neuropharmacology 27, 1061-1065. Yaksh, T. L., Kohl, R. L., and Rudy, T. A. (1977). Eur. J . Pharmucol. 42, 275-284. Yaksh, T. L., Dirksen, R., and Harty, G. J. (1985). Eur. J . Phannacol. 117, 81-88. Yamori, Y. (1976). I n “Regulation of Blood Pressure by the Central Nervous System” (G. Onesti, M. Fernandes, and K. W. Kim, eds.), pp. 65-76. Grune & Stratton, New York. Younkin, S. G., Goodridge, B., Katz, J., Lockett, G., Nafziger, D., Usiak, M. F., and Younkin, L. H. (1986). Fed. Proc., Fed. Am. SOC.Ex$. Biol. 45, 2982-2988.

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DEVELOPMENT OF THE LEECH NERVOUS SYSTEM Gunther S. Stent,* William 6. Kristan, Jr.,t Steven A. Terrence: Kathleen A. French,t and David A. Weisblat* *Department of Molecular and Cell Biology University of California, Berkeley, Berkeley, California 94720 tDepartment of Biology University of California, San Diego La Jolla, California 92093

I. Introduction to the Leech A. Historical Background B. Taxonomy C. Gross Anatomy D. The Leech Nervous System 11. Morphological Development and Staging 111. Behavioral Development IV. Developmental Cell Lineage A. Cell Lineage Tracing B. Genealogical Origins of the Segmental Neurons C. Origin of the Supraesophageal Ganglion D. Transfating V. Myogenesis and Neurogenesis A. Myogenesis B. Gangliogenesis C. Neurochemical Differentiation D. Electrophysiological Differentiation E. Morphological Differentiation F. Interactions between Neurons and Their Peripheral Targets G. Neuron-Neuron Interactions: The Origins of Unpaired Neurons VI. Conclusions References

1. Introduction to the leech

The nervous system presents two of the most challenging questions of contemporary biology: How do networks of neurons generate animal behavior? And how do the neurons and their specific connections arise during the development of an animal from the fertilized egg? The second question cannot be considered independently of the first, because 109 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 33

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the anatomy and function of the adult nervous system represent the endpoint of neural development. That is to say, detailed anatomical and functional knowledge is needed even to ask, let alone answer, wellfocused questions in developmental neurobiology. For gaining such knowledge, the leech with its relatively simple and electrophysiologically highly accessible nervous system is particularly suitable.

A. HISTORICAL BACKGROUND

'The tfierapeutic use of leeches, which had reached the peak of its popularity in the mid-nineteenth century, stimulated basic research on their reproduction, development, and anatomy. For example, leeches were the working material of one of the nineteenth century pioneers of modern experimental embryology, Charles 0. Whitman. In the 1880s, Whitman presented the first analysis of developmental cell lineage, describing the successive cleavages leading from the fertilized leech egg to the early embryo and the subsequent morphogenetic cell movements leading to the juvenile leech. On the basis of his studies, Whitman (1878, 1887) put forward the idea, then quite novel, that each identified cell of the early embryo, and the clone of its descendant cells, play a specific role in development. Despite these highly promising beginnings, embryological interest in leeches declined after the turn of the century. Similarly, the nervous system of the leech was studied by nineteenth century pioneers of modern neuroanatomy, such as Santiago Ramon y Cajal (1904) and Gustav M. Retzius (1891). Neurobiological interest in leeches also declined after the turn of the century, not to be rekindled until the 1960s, when Stephen Kuffler and David Potter first applied modern electrophysiological techniques to study glial cells in the leech nervous system (Kuffler and Potter, 1964). Their work was continued by John G. Nicholls and his students, who ascertained the modalities, receptive fields, and response characteristics of leech sensory neurons (Nicholls and Baylor, 1968), the fields of peripheral action of effector neurons (Stuart, 1970), and the integrative role of connections from the sensory neurons to the motor neurons (Nicholls and Purves, 1970, 1972). These findings, in turn, allowed the identification of specific cells and their synaptic connections responsible for generating not only some simple reflexes, but also some moderately complex integrated motor acts (Stent and Kristan, 1981). Acquiring such detailed knowledge of functional elements within the leech nervous system made it possible to ask specific, focused questions regarding the system's development. For that reason, and in view of the

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classical body of knowledge regarding its embryogenesis (Schleip, 1936; Dawydoff, 1959; Anderson, 1973), the leech seemed to offer considerable promise as an experimental object for developmental neurobiology. Thus studies of leech neurodevelopment began anew in the mid-1970s. This article presents an overview and synthesis of these recent studies; more detailed reviews of specific features of leech development are available '(Weisblat, 1981; Kristan et al., 1984; Weisblat & Shankland, 1985; Stent, 1985; Shankland and Stent, 1986; Weisblat and Astrow, 1989; Levine and Macagno, 1990; French and Kristan, 1991).

B. TAXONOMY Leeches form the class Hirudinea in the phylum Annelida. They are so closely related to earthworms (class Oligochaeta) that, together, leeches and earthworms are assigned to the superclass Clitellata (Anderson, 1973; Sawyer, 1972, 1986). An important difference between leeches and earthworms, however, is the number of body segments. In earthworms that number is variable, owing to continual addition of new segments from a posterior growth zone throughout life, whereas the number of body segments in leeches is constant after early embryogenesis (Harant and GrassC, 1959; Mann, 1962). The constant segment number in leeches may result from their possession of a caudal sucker, a structure that is developmentally incompatible with the type of posterior growth zone found in earthworms (Sawyer, 1981). Constancy of segment number allows a higher degree of specialization in different body regions than is found in earthworms, and leeches differ from earthworms, both anatomically and behaviorally, in ways that make them favorable experimental preparations for studying neurodevelopment. For instance, the earthworm nervous system consists of a diffuse distribution of metameric neurons along the ventral midline, whereas the leech CNS is organized into a chain of discrete segmental ganglia (Bullock and Horridge, 1965). T h e class Hirudinea comprises three orders (Pharyngobdellida, Gnathobdellida, and Rhynchobdellida), of which the latter two have been the object of neurobiological studies. Both of these orders feed by bloodsucking, but gnathobdellids bite the host with toothed, rasping jaws, whereas rhynchobdellids insert a muscular proboscis into the host. Most studies of the adult leech nervous system have been on species in the gnathobdellid order, especially on members of the family Hirudinidae, such as Hirudo medicinalis. In contrast, most neurodevelopmental studies have been on the embryologically more favorable

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FIG. 1. Body plan of the medicinal leech, Hirudo mtdicinalis. (A) Drawing of the dorsal aspect of an adult leech, indicating the location of the ventral nerve cord and its segmental ganglia. The external surface is divided by a series of circumferential grooves; the space between adjacent grooves is an "annulus." The large dots indicate the central annulus of each of the 21 midbody segments, numbered in rostrocaudal order. Most midbody segments contain five annuli, as indicated by the arrows bracketing segment 8, and each central annulus has several sensilla, indicated by small dots. The anterior brain includes a single pair of supraesophageal ganglia and four fused subesophageal ganglia; the posterior brain is composed of seven fused segmental ganglia. There are five pairs of eyes, one pair in each of the segments innervated by the anterior brain and one pair in the first midbody segment. The anterior sucker, containing the mouth, is on the ventral surface, just behind the anterior brain. (B) Drawing o f a transverse cut through the midbody, indicating the locations of the major muscle layers and blood vessels, as well as the nerve cord and the gut. The lateral blood sinuses serve as heart tubes because, unlike the dorsal and ventral blood sinuses, they have muscles in their walls and contract to move blood in the closed circulatory system. Two nerve roots-anterior and posterior-emerge from each ganglion and enter the body wall. The anterior root splits into an anterior and medial

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rhynchobdellid order, especially species in the family Glossiphoniidae: Helobdella, triserialis, Theromyzon rude, and Haementeria ghilianii. Helobdella triserialis is native to North America and feeds on aquatic snails. It reaches an adult length of 1-2 cm and propagates with an egg-to-egg generation time of about 9 weeks (Sawyer, 1972). Theromyzon rude is larger (about 2-4 cm in length) and feeds on the blood of aquatic birds, especially ducks; T. rude has not been successfully cultivated in the laboratory, but can be readily collected from ponds; its generation time in the wild is thought to be 1 year, Haementeria ghiliunii is native to South America and feeds on the blood of mammals. It reaches an adult length of up to 50 cm and has an egg-to-egg generation time of about 10 months in laboratory cultivation (Sawyer et al., 1981). The short generation time, simplicity of cultivation, and hardy embryo of Helobdella make it favorable for developmental studies, but its small size renders it less favorable for neurophysiology. In contrast, the enormous size of Haementeria makes even its embryonic nervous system accessible to recording and injection techniques that require penetrations of single cells, but the long generation time, demanding breeding conditions, and the more fragile embryos present drawbacks in comparison with Helobdella. Theromyzon embryos are as hardy as those of Helobdella and yet are large enough to provide some of the advantages of Haementeria. Fortunately, despite differences in size and habit, the three species are similar in adult body plan and embryonic development so that, for many purposes, the results obtained with one are applicable to the other two.

C. GROSSANATOMY

The tubular body of the leech consists of 32 segments, plus a nonsegmental prostomium (Fig. 1A). The anteriormost four (“head”)segments are fused, forming specialized cephalic structures, including pairs of eyes dorsally (0-9 pairs in different species) and a mouth surrounded by the anterior sucker ventrally. The posteriormost seven (“tail”)segments are also fused, forming the large caudal sucker. Between fused head and tail segments lie 21 unfused midbody segments, designated in rostrocaudal order as M1 to M21. (These ganglia have also been called nerve, and the posterior root splits into a dorsal and posterior nerve. All the major nerves are mixed, containing both sensory and motor axons. Axons run between ganglia via the connectives, consisting of a pair of large connectives and a single smaller connective between adjacent ganglia. (These drawings are variations of ones from Nicholls and Van Essen, 1974.)

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“abdominal” and “segmental” ganglia.) Along the entire length of the body, the skin is subdivided into circumferential rings, or annuli, of which there is a constant, species-specific number per midbody segment. This num’ber is 5 in the case of H i d o , whereas it is 5 on the ventral and 3 on the dorsal aspect of Haementeriu. In all species the number of annuli per segment is reduced in the head and tail segments, as well as in some of the anteriormost and posteriormost midbody segments. T h e central annulus of each segment contains a set of circumferentially distributed sensory organs, or sensilla, which comprise photo- as well as mechanoreceptors (Kretz et al., 1976; Derosa and Friesen, 1981).T h e excretory system of the leech consists of paired, metameric nephridia distributed in a species-specific mode over most of the midbody segments. Each individual nephridium excretes urine via a nephridiopore located on the ventral aspect (Hardnt and GrassC, 1959; Mann, 1962). Segments M 5 and M6 are the reproductive segments. T h e male pore, penis, and vas deferens lie in M5; sperm are produced and stored in metameric testes distributed over several more posterior segments. The paired ovaries lie in M6; eggs are fertilized internally and are laid through the female pore of M6. Below the epidermis lie three layers of muscle fibers (Fig. 1B). The outer layer consists of circumferential “circular” muscle fibers and the inner layFr of longitudinal fibers. T h e intermediate layer is formed by two thin sheets of crossed oblique muscle fibers: the fibers of one sheet lie at an angle of +45” and those of the other sheet lie at an angle of -45” to the longitudinal axis. T h e body of the leech is traversed by a fourth, dorsoventral set of muscle fibers, which insert into the dorsal body wall at one end and into the ventral body wall at the other. The length of individual longitudinal fibers is variable, ranging from about two-thirds of a segment to two segments (Cline, 1986); the lengths of the other muscle fibers in adult animals are not known. All four types of muscle fibers are arranged in discrete parallel fascicles. Each segment contains a fixed number of identifiable fascicles, with a fixed number of each type of muscle fascicle per segment. Contraction of each type of muscle works against the hydrostatic skeleton provided by the fluid-filled leech body tube to effect a characteristic change in body shape: contraction of the circular fibers causes constriction and lengthening, contraction of the longitudinal fibers causes shortening, and contraction of the dorsoventral fibers causes flattening and lengthening. T h e effect of contraction of the oblique fibers depends upon which other types of fibers happen to be contracting. During longitudinal fiber contraction (i.e., in a shortened animal), oblique fiber contraction produces elongation; during circular fiber contraction (i.e., in a fully extended animal), oblique

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fiber contraction produces shortening; when no other fibers are contracted, contraction of the oblique fibers stiffens the body wall at an intermediate body length. A fifth set of muscles, the annulus erectors (Stuart, 1970), is composed of short longitudinal fibers that traverse a single annulus just below the epidermis. Contraction of the erectors raises the annuli, forming a series of sharp ridges that make the epidermis resemble a washboard’s surface. D. THELEECHNERVOUS SYSTEM

1. Ventral Neme Cord T h e leech nervous system reflects the segmental body plan (Fig. 1A). T h e CNS consists of a ventral chain of 32 segmentally iterated ganglia (Mann, 1962). The anteriormost four and posteriormost seven segmental ganglia are fused, constituting rostral and caudal ganglionic masses (or “brains”),respectively. The rostral ganglionic mass, or subesophageal ganglion,’is linked at its anterior end via two circumesophageal connectives to a dorsally situated supraesophageal ganglion. The supraesophageal ganglion is part of the prostomium, and hence, unlike all other ganglia, is not a segmental organ. T h e unfused segmental ganglia are linked via an unpaired, median connective, called “Faivre’s nerve,” and two paired, lateral connective nerves. The connective nerves contain, in addition to interganglionic axons, several longitudinal muscle fibers whose contraction or distension is coordinated with changes in body length caused by the body wall musculature. Each segmental ganglion contains about 200 bilateral pairs of neurons (Macagno, 1980), as well as a few unpaired neurons (Fig. 2). Their cell bodies form an outer cortex around the ventral and lateral aspects of the ganglion. The neurons are monopolar; their processes project initially into a central neuropil, where they make synaptic contacts. From there, the axons of some neurons project to other ganglia via the connective nerves. Sensory and effector neurons send processes to peripheral targets via segmental nerves, whose roots emerge from the lateral edges of the ganglion. From either side of the typical midbody ganglion (Fig. 1B) emerge four main segmental nerves: the anteroanterior (AA), the medioanterior (MA), the dorsoposterior (DP), and the posteroposterior (PP) nerves (Ort et al., 1974). The detailed anatomy of these nerves differs somewhat among various leech species. The segmental nerves arborize extensively in the body wall, but all preserve a main, circumferential nerve trunk. At the dorsal midline, right and left main nerve trunks anastomose to form circumferential nerve rings.

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FIG.2 . Disposition of neuronal cell bodies on the ventral and dorsal surfaces of a segniental ganglion of Hir-udo medicinah. Cell bodies are distributed in a single layer around the cortex of the ganglion. The continuous lines with the ganglion indicate cell packet margins. Filled outlines indicate the approximate position of identified neurons of different function: sensory, motor, modulatory, or interneurons. Also indicated are identified neurons of unknown function, termed “partially characterized,” and as yet uncharacterized neurons. Most of the identified neurons are found in all ganglion, although some are found in specialized regions of the body. [Details of the identity of most of these neurons, a s well as references to their original identification, can be obtained from Muller et al. (1981).]

In each ganglion, the neuronal cell bodies are distributed among six cell packets: a pair of anterolateral packets, a pair of posterolateral packets, and a pair of ventromedial packets. The latter pair lie anteriorly and posteriorly in all but the anteriormost ganglia, where they lie nearly side by side. Each neuronal packet is enveloped by one giant glial cell. In addition, two giant glial cells are associated with the ganglionic neuropil, and additional giant glial cells are present in the interganglionic connective nerves (Coggeshall and Fawcett, 1964; Weisblat et al., 1980b). T h e anatomy of the leech ganglion is sufficiently stereotyped, and its cell bodies are sufficiently accessible to electrophysiological and anatomical analysis, that a substantial fraction of its neurons have been identified (Fig. 2). After characterizing a particular neuron in a particular ganglion of a particular specimen according to morphological and physiological criteria, homologous neurons can usually be found on the other side of that same ganglion, in other ganglia of that same specimen, in the

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ganglia of other specimens of the same species, and even in other leech species, families, or orders (Muller et al., 1981). It is likely that all neurons of the segmental ganglia are identifiable in this sense. Despite this high degree of neural stereotypy, some systematic variations in the number of cells do occur among different segmental ganglia within the same nerve cord and among corresponding ganglia in the nerve cords of different leech species. For instance, in H. medicinal&, the ganglia in the two reproductive body segments M5 and M6 contain nearly twice as many cells as do the other midbody ganglia; in H . ghilianii the corresponding ganglia contain only about 5% more cells than do ganglia in nonreproductive segments (Macagno, 1980).Moreover, some slight variations in the exact number of neurons per ganglion has been found among corresponding ganglia from different individuals of the same species. This “developmental noise” amounts to a variance of about 1% in the total number of neurons per corresponding ganglion, thus placing a conceptual limit on the idealized picture of the segmental ganglion as a fixed set of uniquely identifiable neurons. In fact, in at least one specimen of H . medicinalis, a set of supernumerary neurons corresponding to identified cell types was observed (Kuffler and Muller, 1974).

2 . Identijied Cells About one-quarter of the neurons in the segmental ganglia of H. medicinalis have been identified according to various criteria, including function (Fig. 2). Thus, many cells have been classified as sensory, effector, or interneurons, and their connectivity has been elucidated (Nicholls and Baylor, 1968; Baylor and Nicholls, 1969; Stuart, 1970; Nicholls and Purves, 1972; Lent, 1973; Ort et al., 1974; Thompson and Stent, 1976a,b,c; Friesen et al., 1978; Muller, 1979, 1981; Friesen, 1985; Nusbaum and Kristan, 1986; Lockery and Kristan, 1990b). These surveys have culminated in the description of sensory pathways and of neuronal networks controlling various behaviors, such as body shortening, heartbeat, and swimming (Stent et al., 1978, 1979; Kristan et al., 1988; Friesen, 1989). Despite their considerable phyletic distance from the hirudinid Hirudo, the glossiphoniid species share with Hirudo not only the same general structure of the CNS, but even many of the identified neurons (Kramer and Goldman, 1981). The identified sensory neurons include three types of mechanosensory cells, designated as T (for touch), P (for pressure), and N (for nociception) (Nicholls and Baylor, 1968).Each of these neurons projects its axons from the ganglion to a particular territory of the segmental skin, where its endings form specialized mechanoreceptors that respond specifically to slight (T),moderate (P), or intense (N) deformation of the

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skin (Yau, 1976; Blackshaw and Nicholls, 1979; Blackshaw, 1981a,b). In the typical midbody ganglion there are pairs of T , TD,T,*,P , and P, cells, for which the subscripts v, D, and L designate that the ipsilateral ventral, dorsal, o r lateral skin, respectively, is the principal territory of innervation. T h e Hirudo ganglion has two pairs of N cells, whereas the Haementerza ganglion has only one pair; in both species, each N cell appears to innervate the entire hemilateral skin (Blackshaw, 1981c; Kramer and Goldman, 1981). Ganglia in specialized segments, e.g., the reproductive segments, contain a somewhat different complement of mechanosensory neurons, a pattern that varies from species to species (Johansen et ul., 1984). ‘The identified effector neurons include an ensemble of about two dozen paired excitatory and inhibitory motor neurons, each of which innervates a particular type of muscle in a particular territory within the contralateral segmental body wall (Stuart, 1970; Ort et ul., 1974; Norris and Calabrese, 1990). In addition, there are neurons that have modulatory effects on muscles and other neurons. For instance, the largest neurons in most midbody segments are the pair of Retzius neurons, which, by releasing serotonin, cause mucus release onto the skin surface (Lent, 1973), increase the rate of muscle contraction and relaxation (Mason et ul., 1979; Mason and Kristan, 1982), increase the probability of both swimming (Willard, 1981) and feeding (Lent and Dickinson, 1989), and increase the magnitude of both local bending and shortening movements (Lockery and Kristan, 1991; Wittenberg, 1991). Another pair of neurons, the Leydig cells, large opalescent somata at the posterolateral marginal of each midbody ganglion, influence the strength of the heartbeat (Calabrese and Arbds, 1989) and of the local bending response (Lockery and Kristdn, 1991). T h e identified interneurons include some of the few unpaired cells, the best studied of which is intersegmental interneuron S, whose giant axon courses in the median connective nerve. T h e axons of the single S cell in each ganglion are linked via strong electrical junctions to the axons of the homologous S cells in both the next anterior and the next posterior ganglion, so that an action potential arising in one segmental S cell is rapidly propagated over the entire nerve cord. This chain of electrically linked giant axons forms a fast through-conducting system over the whole length of the leech CNS (Frank et ul., 1975; Magni and Pellegrino, 1978; Muller and Carbonetto, 1979). Mechanosensory neurons are linked to the S cell chain via “coupling interneurons,” a single pair of small neurons that are dye coupled to the S cell in each ganglion (Muller and Scott, 1981).In addition, interneurons have been identified that contribute to the generation of the heartbeat (Thompson and Stent,

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1976a,b,c; Stent et al., 1979; Calabrese et al., 1989), swimming (Friesen et al., 1978; Poon et al., 1978; Weeks, 1982a,b; Friesen, 1985, 1989; Nusbaum and Kristan, 1986; Brodfuehrer and Friesen, 1986), local bending (Lockery and Kristan, 1990b), and shortening (Wittenberg, 1991). Finally, a few neurons have been identified by their morphology, by their electrical properties, o r by their connections to other neurons (Muller et al., 1981; Wadepuhl, 1989), but their behavioral functions remain unknown. There are also identified neurons found only in particular regions of the leech-in the supraesophageal ganglia, for instance, or in the midbody segments devoted to reproduction. These neurons have not been indicated in Fig. 2, and their identifying characteristics will be given as they are discussed in later sections. 3. Neurotransmitters T h e characterization of leech neurons has been extended to the identification of actual or putative neurotransmitters by electrophysiological, pharmacological, autoradiographic, histochemical, and immunohistological techniques. For example, the identified excitatory motor neurons innervating the body wall muscles are cholinergic (Kuffler, 19’78),whereas the corresponding identified inhibitory motor neurons are y-aminobutyric acid (GABA)ergic (Cline, 1986). There may be additional cholinergic and GABAergic neurons in the segmental ganglia, because many other cells contain choline acetyltransferase activity (Sargent, 1977) and a specific cholinesterase activity (Wallace and Gillon, 1982), as do the muscle excitors; other neurons have a high-affinity uptake system for GABA (Cline, 1983, 1986), as do the muscle inhibitors. Furthermore, glutamic acid (the metabolic precursor to GABA), known to serve as a neurotransmitter in other nervous systems, is concentrated in a small number of neurons in the anterior portion of the leech CNS (Brodfuehrer and Cohen, 1990); the release of glutamate is thought to mediate the initiation of swimming behavior. Neurons containing monoamine neurotransmitters are present in the leech as well. Segmentally iterated serotonin-containing neurons, referred to here as “serotonin neurons” (Fig. 3) (Rude, 1969; Stuart et al., 1974), can be identified by several criteria: they emit yellow-green fluorescence after exposure to formaldehyde or glyoxylic acid (Lent, 1982; Stuart et al., 1987), they accumulate the dye neutral red (Stuart et al., 1974), they react with an antiserotonin antibody (Stuart et al., 1982; Jellies et al., 1987), and they selectively accumulate serotonin from the extracellular fluid (Glover and Stuart, 1983). Each segmental ganglion includes three pairs of serotonin neurons: the Retzius (R) cells and two

120

GCNTHER S . Sl‘EN.1 r/ ul.

Frc,. 3. Identification of monoamine-containing cells in a midbody segment of Haemenfe% g/zz/iattzi.This is a summary diagram of monoamine-containing cell types found in midbody segments. based on histofluorescence induced by treatment with glyoxylic acid. The yellow-green fluorescence characteristic of serotonin is seen in nine cell bodies: the giant Retzius (R) cell pairs, the dorsolateral (dls) and ventrolateral (vls) cell pairs, and the paired anteromedial (anis) and unpaired posteromedial (pms) cell pairs. The blue-green fluorescence characteristic of dopamine is seen in three pairs of peripherally located cell bodies ( L D I , LD2, and MD) and their extensive arborizations (not indicated) within the central neuropil. Anterior is up; ganglia in adult leeches are about 500 Frn in width. (Drawing provided by D. K. Stuart.)

other pairs of smaller cells designated as dorsolateral (dls, or cell 2 1) and ventrolateral (vls, or cell 61) neurons. Most ganglia also contain one unpaired posteromedial (pms) neuron (Blair, 1983; Lent et al., 1979; Nusbaum and Kristan, 1986; Leake, 1986; Stuart et al., 1987). In Haementerin and Helobdella, pms is present only in ganglia Ml-M7 (Stuart et ad., 1987), and there is also an anteromedial (ams) pair of serotonin neurons in ganglia Ml-M3. In addition to labeling serotonin neurons, glyoxylic acid treatment induces a blue-green fluorescence in segmentally iterated dopaminecontaining neurons, referred to here as “dopamine neurons” (Fig. 3). One pair, designated as the medial dopamine (MD) neurons, is located just beyond the margin of the ganglion in all midbody segments except M 1. There, and in the head and tail segments, the MD pairs are located within the ganglia (Wallace, 1981). In glossiphoniid species, there are two additional pairs of dopamine neurons, designated as lateral dopamine neurons, o r LD1 and LD2. They lie near the lateral edge of the body wall, within the trunks of the PP and AA segmental nerves, respectively (Blair, 1983; Stuart et al., 1987). T h e dopamine neurons project axons into the CNS, where they arborize profusely throughout the ganglionic neuropil.

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A third biogenic amine, octopamine, has also been identified in the leech nervous system (Webb and Orchard, 1980, 1981). Localization of octopamirie is more problematic than the other two amines because it is not fluorogenic when treated with formaldehyde or glyoxylate, and there is no specific antibody to it. Despite this, radioenzymatic assays have shown that the cell bodies of the Leydig cells account for most of the octopamine in the ganglion (BClanger and Orchard, 1986); octopamine is thought to play a neurohormonal role in modulating activity levels (BClanger and Orchard, 1988). In addition to low-molecular-weight neurotransmitters, several neuropeptide transmitter candidates have been found in the leech nervous system. Neuropeptides related to, or derived from, FMRFamide are probably the best studied of these substances found in the leech. Antibodies to FMRFaniide stain about 40 cells per midbody ganglion, including almost all of the excitatory motor neurons except the excitors to the circular muscles and to the lateral dorsoventral muscles (Kuhlman et al., 1985; Evans and Calabrese, €989; Norris and Calabrese, 1990). Other neurons stained by anti-FMRFamide antibody include a swim-initiating interneuron (cell 204) and an inhibitory motor neuron (cell 101). HPLC analysis of Hirudo nerve cord extracts has revealed the presence of four FMRFamide-like immunoreactive peptides, including FLRFamide, YMRFamide, and, YLRFamide, as well as FMRFamide itself (Evans et al., 1990). The FMRFamide family of peptides modulates neuromuscular activity in the leech, in part by gradually potentiating the response of muscles to acetylcholine (Norris and Calabrese, 1990). Some of the neurons staining with anti-FMRFamide antibody also stain with antibody against the small cardiac peptide (SCP) B (Shankland and Martindale, 1989; Evans and Calabrese, 1989), although chromatographic analyses of Hirudo nerve cord failed to demonstrate the presence of authentic SCP (B. Evans and R. Calabrese, personal communication). Evidence for other leech neuropeptides is limited to immunohistochemical data. These include a proctolin-like immunoreactivity in inhibitory motor neurons (Li and Calabrese, 1985); substance P-like immunoreactivity in a single pair of neurons at the base of the circumesophageal commissures of the head brain (S. Ranganathan, D. K. Stuart, and L. Gleizer, personal communication); red pigmentconcentrating hormone (RPCH)-like immunoreactivity in approximately a dozen neurons in each segmental ganglion, plus others in the head and tail brains, including the supraesophageal ganglion (M. Nusbaum, personal communication); and enkephalin-like immunoreactivity in a pair of neurons in each segmental ganglion (Zipser, 1980).

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4. Sensory a d Motor Fields T h e receptive field of a leech mechanosensory neuron consists of a major field in the ipsilateral skin of the segment in which the ganglion containing the cell body is located and minor fields in the ipsilateral skin of both adjacent segments (Nicholls and Baylor, 1968; Yau, 1976; Blackshaw, 1981a,b,c; Kramer and Goldman, 1981). T h e major field is innervated by an axon that exits from the ganglion containing the cell body via a segmental nerve, and the minor fields are innervated by axon branches that course in the connectives to the adjacent ganglia, where they exit via homologous nerves to the periphery (Fig. 4). In addition, the major and minor sensory fields are composed of distinct subfields, with each subfield being innervated by a separate axon branch coursing within one of the main branches of the segmental nerves. These receptive fields are sufficiently stereotyped to allow a description of their typical characteristics: overall size, major and minor fields, structure, and skin territories of innervation. However, there is significant variation from segment to segment and from specimen to specimen in the number of axonal branches growing out to the skin from homologous cells, and hence in the number of subfields and in the location of the subfield borders (Kramer et al., 1985). Even in cases in which the same number of subfields is present, there can be substantial variation in the size, shape, and position of the subfields making u p the whole field. Thus, although a given mechanosensory neuron seems destined to innervate a specific territory of skin, the actual innervation pattern of that territory is subject to considerable indeterminacy, or “epigenetic noise,” in the developmental program (Waddington, 1957). T h e motor neurons also innervate their targets according to a specific pattern (Stuart, 1970). At the first level of specificity, a particular motor neuron always innervates a single type of muscle fiber. For example, a motor neuron servicing longitudinal fibers does not innervate any circular fibers, either in the course of normal development or after its axon has been cut and directed to a different region of the body wall (Van Essen and Jansen, 1977). At the second level of specificity, a particular motor neuron normally innervates only a subset of the potentially available fibers of the appropriate type (Stuart, 1970). For instance, the excitor designated d innervates longitudinal fibers only near the dorsal midline of the body wall, whereas the excitors 1 and v innervate longitudinal fibers only in the lateral and ventral body wall, respectively. Two further excitors, dl and vl, innervate longitudinal fibers only in the dorsolateral or ventrolateral body wall quadrants, respectively. One excitor,

LEECH NERVOUS SYSTEM DEVELOPMENT

C

123

C

DM

LE

VM

- t - -I--

I

t -1-+PP

I

FIG.4. Receptive field of Pv, a mechanoreceptive neuron responsive to pressure on the ventral surface of the skin of Haementeria ghilianii. The parts of the receptive field innervated by the four different branches of Pv are mapped separately on the schematic representation of three segments of skin on one side of the leech from ventral (VM) to dorsal (DM) midlines. LE marks the lateral edge of the body. Vertical lines represent the margins of the annuli. The central (C) annulus in each segment contains the sensilla (small circles in the central annuli). In this species, the ventral territory in most midbody segments has five annuli and the dorsal territory has three. The complete receptive field is outlined in bold lines, except for the portion that crosses the ventral midline, which has been left out. The horizontally shaded and stippled areas constitute the major subfield: the shaded area is the part innervated by branches of the largest peripheral axon, which leaves the ganglion in the medial (MA) nerve; the stippled area is the part innervated by branches of a smaller axon, which leaves the ganglion by way of the posterior (PP) nerve. The cross-hatched areas are the minor subfields, which are innervated by branches of Pv axons that course through the interganglionic connectives, through the neuropil, and exit via peripheral nerves of the adjacent ganglia. (From Kramer and Goldman, 1981.) The receptive field of the PD neuron in the same segment innervates the dorsal territory; its receptive field overlaps that of Pv across LE. The receptive fields of sensory neurons were originally determined for mechanoreceptors in Hirudo medicinalis (Nicholls and Baylor, 1968);they are very similar to those shown here for Haementeria.

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A. Shortening

B. Local bending

D. Swimming

FIG. 5. Four behaviors whose neuronal basis has been studied in Hirudo medicinalis. (A) In response to a moderate mechanical stimulus anywhere along the body, the animal shortens by contracting all the longitudinal muscles in several segments on either side of the location touched. In response to stronger mechanical stimuli, particularly at the front end, the animal will contract longitudinal muscles in all segments. (B) In response to light or moderate mechanical stimuli, the segment touched will bend away from the stimulus by contracting the longitudinal muscles on the side of the touch and relaxing the longitudinal muscles on the opposite side. This response will occur in the same segment stimulated even when adjacent segments are producing the shortening response. (From Lockery and Kristan, 1990a.) (C) Crawling locomotory behavior is accomplished by an inchwormlike series of movements, using the front (to the right) and rear (to the left) suckers as points of attachment. The extension phase of the step is produced by contraction of circular muscles with the posterior sucker attached to a firm substrate; the contraction phase, in which the body is pulled forward, is accomplished by contraction of the longitudinal muscles with the anterior sucker attached. (From Stern-Tomlinson el al., 1986.) (D) Swimming locomotion is accomplished by a dorsoventral undulation of the body in water with both suckers unattached. Shown are body outlines at about 50-msec intervals, showing a complete cycle of undulation in a leech moving forward from right to left. The body is held extended by a continuous contraction of all the dorsoventral muscles in the body. The crest of each body wave is produced by contraction of the ventral longitudinal muscles in a restricted body region, and the troughs are produced by contraction of the ventral longitudinal muscles

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L, innervates all longitudinal fibers on one side of the segmental body wall. 5. Behavior The behavioral repertoire of leeches ranges from simple reflexes, through locomotion and feeding, to complex mating routines (Gee, 1913; Sawyer, 1981). The simple reflexes elicited by mechanical stimulation of the skin include shortening (Fig. 5A), curling, writhing, and local bending (effected by contraction of the longitudinal muscles on one side of a single segment and distension of the longitudinal muscles on the other side) (Fig. 5B). Reflex implementation varies with the location and intensity of the stimulus (Kristan et al., 1982). Leeches carry out two types of locomotory movements: crawling and swimming. Whereas all known leech species crawl, only some species swim; some species swim as juveniles but stop swimming as they reach adulthood (Sawyer, 1981). Crawling consists of a sequence of stepwise movements (Fig. 5C). At the beginning of each crawling step, the posterior sucker is attached to the substrate. The leech then extends its body forward by contracting the circular muscles and relaxing the longitudinal muscles, while the head searches for a suitable attachment site for the anterior sucker. After the anterior sucker attaches to the substrate, the rear sucker releases its hold and the rear is brought forward by contraction of the longitudinal muscles and relaxation of the circular muscles. Finally, the posterior sucker attaches to the substrate at a more forward location and the anterior sucker is detached. At this point, one crawling step has been completed, and the next can be taken (Stern-Tomlinson et al., 1986). The leech swims by undulating its extended and flattened body in the dorsoventral plane, forming a single wave that travels along the body from head to tail (Fig. 5D) (Gray et al., 1938; Gray, 1968; Kristan et al., 1974a,b). Swimming movements are produced by two types of muscles (von Uexkull, 1905). First, the body flattens by tonically contracting the dorsoventral muscles. Second, rhythmic local contraction and relaxation of longitudinal muscles alternately shorten and lengthen the dorsal and ventral body wall, with the ventral wall relaxed while the dorsal is contracted and vice versa, to form the troughs and crests of the wave. Swimming subserves both food seeking and escape. Starved leeches regionally. Forward progress is made by a coordinated anterior-to-posterior progression of the contraction patterns, such that the crests and troughs move backward along the body; in this manner, the body waves push backward on the water and produce a forward thrust. (From Stent et al., 1978.)

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swim toward the source of water waves created by a host animal (Mann, 1962; Young et al., 1981), a response that appears to be mediated by ciliated mechanoreceptors in the sensilla (Friesen, 198 1 ; Phillips and Friesen, 1982). Moreover, in response to tactile stimulation of its caudal skin, a stationary leech will swim. This response is likely to be mediated by the T and P mechanoreceptor neurons, because electrical stimulation of the skin adequate to activate these neurons usually elicits swimming, particularly when delivered to the rear end of the animal (Kristan et al., 1982). The behavior of leeches depends on their nutritional condition: starved leeches spend much of their time making searching movements and respond to stimuli that sated, quiescent animals ignore (Mann, 1962). These differences in behavior appear to depend upon differences in blood levels of serotonin, because very active leeches have high, and quiescent leeches have low, levels of serotonin in their body fluids (Willard, 198 1). One important source of circulating serotonin is the pair of Retzius cells present in each segmental ganglion. At a high rate of impulse activity, the Retzius cells release enough serotonin into the blood to increase significantly the concentration of circulating serotonin, and hence the frequency and/or duration of swimming episodes. Impulse activity in the two smaller, paired serotonin neurons (cells 21 and 61) also stimulates swimming. In contrast to the neurohormonal release of serotonin into the general circulation by the Retzius cell, however, the smaller cells make direct synaptic contact with, and provide strong excitation to, one of the interneurons that form part of the segmental neuronal circuit that generates the swim motor pattern (Kristan and Nusbaum, 1983; Nusbaum and Kristan, 1986). Serotonin also appears to play a role in the control of feeding (Lent et al., 1988; Lent and Dickinson, 1989). For instance, activity in serotonin neurons of the subesophageal ganglia and the anterior midbody ganglia evokes such feeding-related activities as pharyngeal pumping and salivation. Serotonin also intensifies simple reflexes, such as local bending (Lockery and Kristan, 1991) and shortening (Wittenberg, 1991), so that it may serve as a general enhancer of leech motor routines. Leeches can modify their behavior as a result of environmental contingencies. For instance, the body-shortening reflex is subject to classical conditioning (Henderson and Strong, 1972; Sahley and Ready, 1988), habituation, and sensitization (Belardetti et al., 1982; Boulis and Sahley, 1988). Other behaviors, such as local bending (Lockery and Kristan, 1991), crawling (Sahley and Ready, 1988), swimming (Debski and Friesen, 1986), and food preference (Karrer and Sahley, 1988) also exhibit various forms of conditioning.

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6. Identijied Circuits The simplicity, stereotypy, and experimental accessibility of the leech nervous system offers the possibility that all of the neurons participating in the generation of any leech behavior can be identified and their synaptic and humoral interactions characterized. This promise is strengthened by the possibility of eliciting many motor acts from semiintact preparations in which the nervous system has been exposed (Gray et al., 1938; Kristan, 1982; Kristan et al., 1974a, 1988; Thompson and Stent, 1976a,b,c; Lockery and Kristan, 1990a,b; Baader and Kristan, 1990; Wittenberg, 1991). In some cases, the motor neuron activity pattern characteristic of a behavioral act can be evoked in the isolated nerve cord, or even in parts of it (Kristan and Calabrese, 1976; Thompson and Stent, 1976a; Lockery and Kristan, 1990a; Wittenberg, 1991).To various degrees of completeness, neuronal circuits have been identified for four behaviors; local bending (Kristan, 1982; Lockery et al., 1989; Lockery and Kristan, 1990a,b), body shortening (Wittenberg, 199l), swimming (Stent et al., 1978; Friesen and Stent, 1977; Stent and Kristan, 1981; Kristan and Weeks, 1983; Friesen, 1989), and heartbeat (Stent et al., 1979; Stent and Kristan, 1981; Calabrese and Peterson, 1983; Calabrese and Arbas, 1989; Calabrese et al., 1989). These characterizations of neuronal circuits provide the points of departure for ascertaining the physiological, morphological, and molecular mechanisms responsible for the development of behavior.

II. Morphological Development and Staging

In all leeches, fertilization is internal and embryonic development begins as soon as the eggs are laid. Helobdella, Theromyzon, and Haementeria lay yolk-rich eggs about 0.4,0.8, and 2 mm in diameter, respectively. The eggs are laid in clutches, enclosed in transparent, soft-walled, salinefilled cocoons, which remain attached to the ventral body wall of the brooding parent for most of embryonic development. However, for experimental purposes, the eggs can be removed from the cocoon at any stage of development and cultured to maturity in saline, the composition of which resembles that of the cocoon fluid. The egg yolk provides the energy and organic matter needed for development, which proceeds under conditions of nearly constant volume. Upon exhaustion of the yolk, by which time the volume of the embryo is still not much larger than that of the egg, the juvenile leech takes its first meal from a host

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A

B

Vent ra I midline

Stage 7 (early)

Germinal plate Germinal band

Stage 7 ( m i d d l e )

m

Stage 8 ( l a t e )

Stage 10

W

Stage 8 (early)

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animal. Subsequent postembryonic growth and maturation of the juvenile leech occur by way of increases in both cell size and cell number (Weisblat, 1981). The embryonic development of glossiphoniid leeches has been divided into l l stages, beginning with egg deposition and extending up to the point at which the juvenile leech is ready for its first meal 3 (Helobdella) to 6 (Theromyzon and Haementeria) weeks later (Fernandez, 1980; Weisblat et al., 1980b; Stent et al., 1982; Bissen and Weisblat, 1989); some stages have been further divided into substages. The definition of the stages is based on morphological criteria discernible in living embryos; the staging scheme and cell nomenclature presented here include modifications and additions that reflect an increasing knowledge of the details of development. Figure 6 shows a schematic representation of key stages in the development of Helobdella. The criteria for the beginning and end of each stage are given in Table I. These stages are equally applicable to Theromyzon and Haementem'a. When first laid, the glossiphoniid leech egg is filled throughout with colored yolk and enclosed in a clear vitelline membrane. T h e first embryonic axis is marked initially by the production of two polar bodies at the animal pole early in stage 1 (Fig. 6A). Later, as the egg approaches its first cleavage, this axis is reinforced by domains of yolk-free cytoplasm, or teloplasm, which form at the animal and vegetal poles. The first cleavage, yielding blastomeres AB and CD, is meridional and establishes the second embryonic axis (stage 2). The second cleavage is also merid-

FIG.6. Embryonic development in glossiphoniid leeches. (A) A lineage tree summarizing the cell divisions leading to formation of polar bodies (pb), macromeres (capital letters A-D and combinations), micromeres (lowercase letters with primes), teloblasts (capital letters M, N, 0, P, and Q), and blast cells (lowercase letters without primes). Each cell division is indicated by a horizontal line, and the spacing between horizontal lines is proportional to the approximate times between subsequent cell divisions. The fact that there are bilaterally symmetric lineages after the cleavages of cells DM" and DNOPQ"' is denoted by broken horizontal lines. Embryonic stages are indicated at left. (B) Schematic representation of embryonic stages, showing the arrangement of teloblasts and their primary blast cell bandlets within the germinal band and germinal plate in early stage 8 (left) and views of whole embryos at selected stages (right). In the early stage 7 embryo, teloblasts have begun to produce blast cells in contact with the micromere cap; by midstage 7, the bandlets have formed recognizable germinal bands. At early stage 8, the heart-shaped germinal bands have begun to coalesce along the future ventral midline; this process leads to the formation of the germinal plate, which is complete by the end of stage 8. By stage 10, segmental tissues, including the segmental ganglia of the ventral nerve cord (black), are well differentiated. Late stage 8 and stage 10 embryos are as viewed from the ventral aspect; all other stages are as viewed from the animal pole. (From Bissen and Weisblat, 1987.)

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TABLE I LEECHEMBRYOGENESIS

STAGES OF (;LOSSIPHONIID

Stage

Stage name

1

Uncleaved egg

2 3 4a

T w o cells

4b

Macromere quintet

4c

Mesoteloblast formation

5

Ectoteloblast precursor

6a

N teloblast formation

6b

Q teloblast formation

7 8

Germinal band formation Germinal band coalescence

9

Segmentation

10

Body closure

11

Yolk exhaustion

Juvenile

Four cells Micromere quartet

Beginning

Egg laying Onset of first cleavage Onset of second cleavage Onset micromere formation (cleavage of D to form D’ + d’) Onset of D’ macromere cleavage to form DM + DNOPQ Onset of cleavage of cell DM” to form left and right M teloblasts Onset of cleavage of cell DNOPQ“‘ to form left and right NOPQ proteloblasts Onset of cleavage of cell NOPQ“ to form N and OPQ Onset of cleavage of cell OPQ” to form OP and Q Completion of cleavage of cell OP Onset of coalescence of left and right germinal bands Completion of germinal band coalescence Appearance of coelomic space in the 32nd somite Completion of fusion of the lateral edges of the germinal plate along the dorsal midline Exhaustion of the yolk in the embryonic gut and first feeding

ional; it divides the egg into four blastomeres, A, B, C, and D (stage 3), and D receives most of the teloplasm. The third cleavage is highly unequal, producing four vegetal macromeres A‘, B’, C’, and D‘ and four micromeres a’, b’, c’, and d’ (stage 4a); D’ then cleaves equatorially to yield cells designated DNOPQ and DM, while cells A’, B’, and C’ undergo another round of micromere production, yielding macromeres A , B”, and C and micromeres a”, b”, and c’’ (stage 4b). At this stage, according to the classical germ layer interpretation, macromeres A , B”, and C” constitute presumptive endoderm, and DNOPQ and DM constitute presumptive ectoderm and mesoderm, respectively (Whitman, 1887). Subsequently (stages 5 and 6), quadrants A, B, and C each undergo one more round of micromere production, whereas the D quadrant cells undergo a series of cleavages to generate five bilateral pairs of large,

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yolky stem cells called M, N, O/P, O/P, and Q teloblasts, as well as additional micromeres. These cleavages are presented schematically in Fig. 6A. Each teloblast carries out a series of 40-100 highly unequal cleavages, producing a bandlet of small primary blast cells (stage 7). T h e bandlets produced by the five teloblasts on either side of the midline rise to the surface and merge to form two ridges of cells, the right and left germinal bands. Between the germinal bands lies a cluster of cells, the “micromere cap,” derived mainly from the micromeres. The blast cells and their bandlets that are produced by the M, N, and Q teloblasts are designated as m, n, and q, respectively. In the germinal bands, the bandlets produced by the two O/P sister teloblasts lie between the n and q bandlets, with the bandlet lying next to the n bandlet being designated as o (and its O/P teloblast of origin being designated as “generative 0 teloblast”) and the bandlet lying next to the q bandlet being designated as p (and its O/P teloblast of origin being designated as “generative P teloblast”) (Shankland and Weisblat, 1984). In Theromyzon, the identities of the generative 0 and P teloblasts are apparent from the positions of the teloblasts themselves, which are simply designated 0 and P (Keleher and Stent, 1990). At this point, the ectodermal bandlets lie in mediolateral order q, p, 0, and n, whereas the mesodermal m bandlet lies under the four ectodermal bandlets (Fig. 6B). As the result of morphogenetic processes that are only poorly understood at present, the left and right germ bands come to join one another at their distal (future anterior) ends at the site of the future head (Fernandez and Olea, 1982). The areas between the bands, and the bands themselves, are covered by a simple squamous epithelium derived from the micromere cap. With ongoing blast cell production, the midportions of the bands move apart over the surface of the embryo, and then gradually, during stage 8, meet and coalesce zipperlike, from the future head rearward along the ventral midline, forming a sheet of cells called the germinal plate (Fig. 7A). The circumferential movement of the germinal bands prior to their coalescence on the ventral midline reverses the mediolateral order of the bandlets, so that the left and right n bandlets come into apposition at the ventral midline. As the germinal bands move over the surface of the embryo, the micromere-derived epithelium expands with them, eventually covering the entire surface of the embryo. Over the part of the embryonic surface not covered by the germinal plate, a superficial, simple squamous epithelium and an underlying layer of circumferentially oriented muscles combine to form a two-layered tissue, the provisional epithelium, which does not contribute to the eventual juvenile leech.

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With the proliferation of cells of the germinal plate to form the body tissues, the plate gradually thickens and expands over the surface of the embryo into dorsal territory. Even before the germinal bands have started to coalesce, the left and right mesodermal bandlets become partitioned into a series of discrete blocks of cells corresponding to hemisomites. During stage 8 and continuing through stage 9, the coelom arises as a cavity within each somite, so that the germinal plate becomes partitioned along its length into a series of tissue blocks, each separated from the others by transverse septa (Fig. 7). Each block corresponds to a future body segment. Segmentation starts at the front and progresses rearward; by the time the ganglion of the hindmost segment has formed (at the end of stage 9), the expanding germinal plate covers about onethird of the ventral surface. The embryo hatches from the vitelline membrane during stage 9. In stage 10, right and left leading edges of the expanding germinal plate meet and coalesce on the dorsal midline, closing the leech body. During this expansion of the germinal plate, the provisional integument retracts before, o r is pushed back by, the leading edges of the plate. By the time that the leading edges of the plate have coalesced on the dorsal midline, the crumpled remainder of the provisional integument has disappeared from the embryonic surface. Meanwhile, formation of the gut is underway. It first appears as a cylinder (filled with yolk provided by macromeres A, B, and C and the remnants of the teloblasts) and then becomes segmented by annular constrictions, which probably correspond to the segmental septa. These constrictions give rise to paired gut lobes, o r caeca, in register with the midbody segments. Gut segmentation is completed at body closure (at the end of stage 10); the embryo now has the general shape of the adult leech. The final steps of morphological development, including maturation of the posterior sucker, occur during stage 11. When the yolk in the gut is exhausted (at the end of stage 1 l), the juvenile leech is ready for its first meal. This description of embryogenesis up to the formation of the germinal plate applies to glossiphoniid leeches in general, such as Tfwromyzon, Helobdellu, and Huementeria, but not to the hirudinids, such as Hzrudo. Hirudo eggs are much smaller (about 0.1 mm in diameter). They contain little yolk and are deposited in a sealed cocoon that contains an albuminous fluid. T h e initial stages of embryogenesis produce a sac, inappropriately called a “Iarva,” complete with a mouth that ingests the albumin, providing an exogenous source of organic matter and energy for subsequent development. T h e embryo forms on the surface of this sac and rapidly increases in size. As in glossiphoniid leeches, embryogenesis in Hirudo proceeds via five pairs of small teloblasts that

B

A Anterior

Mouu7 ReproductiveStructures

Protonephridia

Larval Sac

Caudal Swker

-1

mm

FIG. 7. Drawings of leech embryos at the stage when ganglia are formed and neurons are growing their processes. (A) Drawing of Haementeria ghzlianii embryo at midstage 9. (From Kuwada and Kramer, 1983.) (B) Drawing of H i d o medicinalis embryo at a comparable stage, E12. The protonephridia and larval sac are nonembryonic tissues that are likely not to be retained in the adult leech. The mouth is at the anterior end. (Drawing provided by J. Jellies.) Embryos vary in size at this stage over a twofold range, hence the calibration is approximate.

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produce bandlets of primary blast cells, germinal bands, and a germinal plate. In the hirudinid embryo, germinal bands form on the future ventral (rather than on the future dorsal) surface and coalesce directly, without undergoing the circumferential migration characteristic of the glossiphoniids (Schleip 1936). Subsequent development follows much the same course in the hirudinids (Fig. 7B) as in the glossiphoniids (Schleip, 1936; Fernandez and Stent, 1983). N o staging system has yet been devised for the embryonic development of the hirudinids such as Hirudo. Instead, the developmental progress of Hirudo embryos is reported in days of development, from egg deposition (EO) to completion of the posterior sucker and maturation of the juvenile leech about a month later (E30). Formation of the posteriormost ganglion of H i d o is complete at about E10, and body closure is complete at about E20, corresponding to the end of stages 9 and 10, respectively, in glossiphoniid embryos.

111. Behavioral Development

The highly regular morphological development of the leech embryo is accompanied by similarly regular behavioral development. T h e behavioral repertoire of a given species of leech emerges in a stereotyped sequence. By the end of embryogenesis (stage 1 1 in glossiphoniids and about E30 in hirudinids), most adult behavioral routines have appeared, except those pertaining to reproduction. A schematic summary of the behavioral development of Haementena is presented in Fig. 8. During the first seven stages, i.e., for many days after the fertilized Haementeria egg is laid, there are no movements, either spontaneous or evocable, in the developing embryo. But as the germinal plate nears completion, peristaltic movements begin. These movements consist of longitudinal waves of circumferential constrictions that usually start at one end of the embryo and take 5-10 sec to reach the other end. Although they arise spontaneously and at irregular intervals, they can also be initiated by gentle mechanical stimulation at any site along the embryo. T h e constrictions are produced by the circumferential muscle cells of the provisional integument. These cells (which are not precursors of the definitive circular muscle fibers) disappear by the time body closure is complete, when peristaltic movements have ceased as well (A. P. Kramer, unpublished observations). T h e peristaltic movement leads to hatching of the embryo from the vitelline membrane. In addition, peristalsis may also circulate fluid in the developing embryo. T h e peristaltic

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rhythm is almost certainly myogenic in origin, because peristalsis begins before the embryonic nervous system becomes functional, and it can persist even after surgical removal of the germinal plate and its developing nervous system. T h e body-shortening reflex appears when body closure is almost complete. At first this reflex is evoked only by mechanical stimuli applied to the very front end of the embryo, and shortening occurs only in the anterior third of the animal. Over the next few days, the receptive field for the stimulus enlarges to cover the anterior half of the animal, and the extent of the shortening response enlarges to encompass most of the body. Within a day after shortening appears, another reflex, namely local body bending (Kristan, 1982), emerges. A clutch of embryos remains enclosed within its cocoon until stage 10, when the embryos emerge and attach themselves directly to the venter of their brooding parent by means of a sticky exudate, localized at the anteroventral end of the embryo. Early in stage 1 1, first the posterior and then the anterior suckers develop and begin to function, allowing the nearly mature embryo to control its attachment to solid substrates. By midstage 11, the embryos are able to use the suckers for wellcoordinated crawling. Before this, during a precrawling phase, the embryos execute a progression of movements that appear to be increasingly complex elements of crawling behavior (A. P. Kramer, unpublished observations). Toward the end of stage 11, the Haementeria embryo acquires the ability to swim, having undergone a preswimming phase during which it produces progressively more swimminglike movements. Is this gradual improvement in swimming performance the result of an autonomous developmental process, or does it depend upon practice? To answer this question, all serotonin neurons of the developing nervous system were ablated during stage 10, i.e., long before the onset of swimming, by injecting the embryo with 5,7-dihydroxytryptamine,a toxic analog of serotonin, (Glover and Kramer, 1982). Because (as will be discussed in more detail later) there is no regulative restoration of ablated neurons in leech development, embryos thus treated develop into juveniles whose nervous system is devoid of serotonin neurons. Despite their lack of what might seem to be a vital neurologic component, the treated leeches are morphologically and behaviorally quite normal, although they are lethargic and do not swim. Such leeches, which have never previously executed a body wave, start to swim as soon as they are either injected with serotonin o r are simply put into water containing serotonin. I t appears, therefore, that the capacity to generate the basic swimming rhythm, and hence the underlying neural circuit, is as-

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sembled via an autonomous developmental process that does not require practice. Even after they are able to swim, Haementeria embryos remain attached to the venter of the parent. Finally, when their store of yolk is exhausted, the embryos detach from their parent in the presence of a potential host animal. If the host has a sufficiently thin skin, the embryo inserts its proboscis through the skin and, by means of pumping movements of its pharyngeal muscles, ingests as much as 15 times its own body weight of blood. With this first feeding, the embryo advances to the status of juvenile leech. T h e behavioral development of hirudinids, which spend their entire embryonic development bathed in an albuminous fluid within a cocoon, is on the whole quite similar to that of the glossiphoniids (Fernandez and Stent, 1983). However, in the hirudinids the order of maturation of the two locomotory behaviors is reversed: they swim before they crawl, and both behaviors appear in embryos at a time when they would normally still be within their cocoon. If the embryos are removed from the cocoon and dropped into water the day after body closure becomes complete (corresponding to early stage 1 1 in glossiphoniids), they make primitive swimming movements, which over several days mature into the adultlike swimming rhythm (Reynolds and Kristan, 1989). Crawling movements and the competent use of the suckers first appear about halfway between body closure and normal emergence from the cocoon. Interestingly, the order in which the two locomotory behaviors appear matches the needs of leech embryos: the rhynchobdellids must use their suckers to attach to the parental venter during the late stages of embryogenesis and need to swim only upon the completion of development, when they must get to a host for their first feeding; the gnathobdellids, by contrast, may swim in the cocoon fluid, and need to crawl only after they emerge, to aid in finding a site for taking blood from their first host.

IV. Developmental Cell Lineage

A. CELLLINEAGE TRACING

To begin to address the fundamental question of which differentiated properties of an adult cell depend on the cell’s line of descent from the fertilized egg and which depend on interactions with other cells, it is necessary to ascertain exact developmental pedigrees. Whitman ( 1 878,

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1887) carried out the pioneering studies on developmental cell lineage in leech embryogenesis over a century ago. His method for ascertaining cell lineages was to observe living embryos under a microscope and keep track of successive cleavages. Cell lineage analyses were later extended to embryos of other species, not only by direct observation, but also by use of such techniques as selective cell ablation, application of extracellular marker particles, and production of chimeric and genetic mosaics (Wilson, 1892; Sturtevant, 1929; Tarkowski, 1961; Mintz, 1965; Stern, 1968; Garcia-Bellido and Merriam, 1969; Le Douarin, 1973; Sulston and Horvitz, 1977, 1981; Deppe et al., 1978; Sulston et al., 1983). More recently, Whitman’s century-old cell lineage studies on leech embryos have been refined and extended by the introduction of microinjected cell lineage tracers (Weisblat et al., 1978, 1980a; Gimlich and Braun, 1985; Stuart al., 1989) to establish the lines of descent of identified cells in the leech nervous system. In this procedure, a histologically detectable tracer molecule is injected directly into an identified cell of an embryo early in development. At a later embryonic stage, the cellular distribution of the tracer is observed. Cells containing the tracer are inferred to have descended from the originally injected cell. For this method to be useful the tracer molecule has to meet three conditions: (1) it must permit embryonic development to continue normally after it is injected, (2) it must remain intact and not be diluted too much during increases in cell size and number within the developing embryo, and (3) it must not pass through junctions linking embryonic cells, so that it is confined exclusively to descendants of the injected cell. (Because early hirudinid embryos have very small cells, microinjection has been used only in glossiphoniid embryos.) Horseradish peroxidase (HRP) was the first molecule to be employed as an intracellular lineage tracer (Weisblat et al., 1978). Soon thereafter covalently linked composites of large carrier molecules and the fluorescent dyes rhodarnine o r fluorescein came into use (Weisblat et al., 1980a). Presently, the most widely used carrier molecules are dextrans; the rhodamine-dextran tracer is designated RDX, and the fluorescein-dextran tracer is designated FDX. To obtain a fluorescent lineage tracer that binds to tissues after histological fixation, dextrans with linked lysine residues are used as carrier molecules (Gimlich and Braun, 1985; Stuart et al., 1989). These fixable composites of rhodamine o r fluorescein and lysinated dextran are designated RDA or FDA. In addition to its utility for tracing cell lineages, a fluorescein-labeled tracer can serve as a specific photosensitizer (Shankland, 1984). Upon illuminating an FDX-labeled cell at a wavelength of about 490 nm, some of the excited fluorescein fluorophores are quenched by transferring the

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absorbed energy to oxygen molecules in solution, converting them to the highly reactive singlet state. These reactive oxygen molecules, in turn, cause a generalized oxidation of cell constituents, killing the illuminated labeled cell (Miller and Selverston, 1979; Braun and Stent, 1989b). Cells are largely transparent to light at 490 nm unless they contain fluorescein, so injecting an embryonic progenitor cell with FDX makes it possible to photoablate its progeny selectively, even if they have become intermingled with cells from other lines of descent. Moreover, because the photosensitizer is also a lineage tracer, it automatically allows direct visual identification of cells to be ablated, as well as direct visualization of their normal fates in unirradiated control embryos.

B. GENEALOGICAL ORIGINS

OF THE

SEGMENTAL NEURONS

Because the paired n blast cell bandlets straddle the ventral midline of the germinal plate, Whitman (1887) and his contemporaries (Bergh, 1885) inferred that the cells of the ventral nerve cord are derived from the N teloblasts. However, using cell lineage tracers has shown that the leech nerve cord is, in fact, derived from all five teloblasts (Weisblat et al., 1984; Kramer and Weisblat, 1985; Weisblat and Shankland, 1985; Torrence and Stuart, 1986), as Apathy (1889) had suggested. The tracerlabeled descendants of each injected teloblast form a distinct pattern, which is repeated from segment to segment and is the same in every embryo in which a particular teloblast has been injected. Figure 9 illustrates the cells that each of the five teloblasts contributes to a midbody segment. Note that the labeled structures lie on the same side as that of the injected teloblast, except for axons projected contralaterally by labeled neuronal cell bodies. Hence in the course of neurogenesis there is no appreciable migration of neuronal precursor cells across the ventral midline. Detailed scrutiny of lineage-tracer-labeled stage 10 embryos shows that each of the four ectodermal precursor teloblasts-N, 0, P, and (2contributes some cells to the CNS, some to the peripheral nervous system, and some to the epidermis. The mesodermal precursor teloblast, M, contributes three to four pairs of neurons to each segmental ganglion, but its main contribution is to mesenchyme, nephridia, and muscle fibers, tissues to which embryologists traditionally assign a mesodermal origin. Thus, each individual teloblast generates progeny that are found in several types of tissue. Descendants of the n blast cell bandlet are found exclusively within the segmental ganglia, except for two or three peripheral neurons

GANGLION

t

VENTRAL MIDLiNE

FIG.9. Contributions of the ectodernial teloblasts and bandlets to a midbody segment. Each panel illustrates the pattern of tracer-labeled cells observed in one segment of'a stage 10 embryo of 7%rromyzon rude following injection of a lineage tracer into one ectodernial teloblast at stage 6 or 7. The right edge of the figure corresponds to the lateral margin of the germinal plate, and thus to the future dorsal midline. Anterior is uppermost. Major labeled axonal tracts are shown, but no attempt has been made to render detailed neuronal projection patterns within the CNS. LDI and LD2 designate identified dopaminecontaining neurons, as in Fig. 3. Names of other neurons (e.g., 921) include a letter indicating the bandlet of origin, the letter z to symbolize the (largely unknown) intermediate cell lineage, and a number. Labeled squamous epidermal cells are indicated by light stippling; epidermal rell florets (CF I -6), are more densely stippled. Top ganglion: Contrihution of the q handlet; AV, anterovenrral cluster of central neurons; CG, connective glioblast; MA, cluster of peripheral neurons along the MA nerve. This cluster includes the dopamine-containing neuron MD. Second ganglion: Contribution of the p bandlet. Third ganglion: Contribution of the o handlet; AD, anterodorsal cluster of central neurons; PV, posteroventral cluster of central neurons. Fourth ganglion: Contribution of the n bandlet; fine, labeled processes fill most of the contralateral neuropil (marked by an asterisk and outlined by a dashed line). (From Torrence and Stuart, 1986.)

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(depending on the species) and a few ventral epidermal cells. Descendants of the 0, p, and q blast cell bandlets contribute substantially fewer cells to the segmental ganglia and correspondingly more cells to the peripheral nervous system and the epidermis, including epidermal specializations called cell florets (Fig. 9). The stereotyped patterns of peripheral neurons that are derived from each blast cell bandlet indicate that the peripheral nervous system also consists of individually identifiable neurons, whose unique identity can be determined based on their position in the body wall and their teloblast of origin. The topography of the contribution each blast cell makes to the segmental epidermis is generally consistent with its relative position within the germinal plate. Thus, dorsal epidermis derives from the q bandlet, ventral epidermis derives mainly from the mediolateral o and p bandlets, and a few epidermal cells on the ventral midline derive from the n bandlet (Weisblat et al., 1980b). The ganglion cells contributed by any blast cell bandlet form discrete and coherent cell domains; they are not randomly distributed or uniformly mixed with those contributed by other teloblasts (Fig. 10). For instance, the n bandlet contributes two transverse slabs of cells in the anterior and posterior regions of the ipsilateral hemiganglion and a longitudinal band of cells adjacent to the midline on the ventral aspect of the ganglion. The cell domains derived from the other bandlets display similarly compact and stereotyped topographies. In fact, these other bandlets contribute smaller numbers of neurons, allowing some of their progeny to be individually recognized. For example, some of the progeny of the o bandlet lie in distinct anterodorsal (AD) and posteroventral (PV) neuron clusters, and some p bandlet progeny form a transverse, wedge-shaped cluster of neurons in the middle of the ventral aspect of the ganglion, including a single neuron just beyond the tip of the wedge. These regular, segmentally iterated patterns of neuronal descent indicate that the developmental cell lineages derived from each blast cell bandlet correspond to four identifiable Kinship groups (Stent et al., 1982; Weisblat et al., 1980b; Weisblat and Shankland, 1985) designated M, N, 0, P, and Q, in accord with their teloblasts of origin. The stereotyped location of each set of teloblast descendants within the embryonic ganglia suggests that each identified neuron and glial cell normally arises from a particular blast cell bandlet. This supposition has been confirmed by surveying the kinship groups in studies combining fluorescent lineage tracers with electrophysiological, anatomical, and histochemical techniques used for identifying individual neurons (Blair, 1983; Kramer and Weisblat, 1985; Stuart et al., 1987). Some results of this survey are summarized in Fig. 10. Do the members of a given neuronal kinship group share a unique

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Dorsal

AA MA

P

AA MA

AA MA

P

P

W

W

Q Ventrol

mcG

AA MA

P

FIG. 10. Schematic representation of the five kinship groups in a typical midbody ganglion of HaPmentrria glrilZa7& The N and 0 kinship groups occupy both dorsal and ventral aspects of the ganglion; P and Q kinship groups are confined to the ventral aspect; the M kinship group is divided between the dorsal aspect of the interganglionic connective (in the case of the muscle cells) and the center of the hemiganglion, midway between dorsal and ventral aspects (in the case of the mz neurons). The connective tracts traverse the dorsal aspect of the ganglion. Boundaries of cell packets, each of which is associated with a packet glial cell, are indicated by dashed lines. The cells in each kinship group are indicated as follows: large cross-hatched regions in N and 0 are clusters of uncounted cells; cross-hatched circles in M,P, and Q are cell bodies of single, unidentified neurons; solid circles with labels are cell bodies of identified neurons; open circles enclosing small solid cirrles denote glial cells. The neuropilar glial cell body is at the ventral edge o f t h e neuropil. The clusters of N-derived cells in the dorsal anterior region of the ganglion are ventral to the dorsal anterior cluster of 0-derived cells. Cell abbreviations are as follows: MCM and LCM, medial and lateral connective muscle cells, respectively; NG, neuropil glia; ALG. anterior lateral giant; K, Retzius neuron; AE, annulus erector motor neuron; P h i l , posteroniedial neuron; AL2 and AL3, anterolateral neurons; LPG and MPG, lateral

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set of properties that set them apart from the members of the other kinship groups, such as functional category (glial cell or sensory neuron, motor neuron, or interneuron) or type of neurotransmitter? The provisional answer to this question must be “no,” because current data (Fig. 10) reveal no obvious kinship group-specific neuronal properties (except that all serotonin neurons belong to the N kinship group and that the 0 and P kinship groups contain some apparently homologous mechanosensory, glial, and dopamine cells). For example, each of the four ectodermal kinship groups includes one or more glial cells; the N, 0, and P kinship groups contain mechanosensory neurons; and the 0, P, and Q kinship groups contain dopamine neurons. How many primary blast cells derived from each teloblast contribute to founding one hemisegmental primordium? The answer to this question turned out to be somewhat complex, but it provided insights into the process of segmentation in leeches. An upper limit of three blast cells has been suggested, because the total number of blast cells produced per teloblast is less than three times the total number of segments (Fernandez and Stent, 1980). The actual number of founder blast cells contributed by each bandlet was estimated by two different methods using lineage tracers: an indirect method, termed the “label boundary method” (Fig. 11) (Weisblat et al., 1984; Weisblat and Shankland, 1985; Bissen and Weisblat, 1987), and a direct method, termed the “double-label method” (Zackson, 1984). Both methods led to the same answer, namely that a single primary blast cell in each of the m, 0, and p bandlets contributes the entire hemisegmental complement of its kinship group, whereas two successively born primary blast cells in the n and q bandlets each contribute a specific subset to the hemisegmental complements of their respective kinship groups. The first-born (i.e., more anterior) of these two blast cells in each hemisegment are called ns and q,, and the younger (i.e., more posterior) cells are called n, and qp A beginning has been made in extending this lineage analysis beyond identifying which primary blast cell gives rise to the roughly 50-100 definitive differentiated cells in each hemisegmental kinship group. First, the initial division patterns of all seven primary hemisegmental founder blast cells were ascertained by observing the caudorostral disposition of second-, third-, and higher-order blast cells in m, n, 0, p, or q and medial packet glia, respectively; PD and Pv, dorsal and ventral pressure-sensitive mechanosensory neurons; the neurons of the P teloblast (pz1-4) and q teloblast (921-3) lineages; TD, TL, and Tv, dorsal, lateral, and ventral touch-sensitive mechanosensory neurons, respectively; N, nocioceptive mechanosensory neuron. The major peripheral nerves (AA, MA, and P) are shown. Anterior is up. (From Kramer and Weisblat, 1985.)

m

a

k-JLA.2 144

FIG. 1 1. Distribution of progeny from individual primary o or p blast cells, ascertained by the labeled boundary method. Neurons labeled with tracer are shown as solid black; labeled epidermal cells are stippled. Segmental ganglia are illustrated as oval outlines, and the ventral midline is indicated by a dashed line through the ganglia. Anterior is up. (A) Three segments from a stage 10 embryo of Helobdelh triserialis in which the generative 0 teloblast was injected with lineage tracer after it had produced some unlabeled blast cells. In the anteriomost segment containing labeled cells (the top segment illustrated), all 0 pattern elements are present but only a subset are labeled. Thus, this segment was populated in part by progeny of the last unlabeled blast cell produced before the teloblast was injected and in part by progeny of the first labeled blast cell. Because the same subset of pattern elements was found to be labeled regardless of where the boundary fell along the longitudinal axis of the embryo, all o blast cells must produce the same complement of progeny. (B) Two segments in an embryo in which the generative P teloblast was injected. As in the 0 boundary segment, only a subset of pattern elements is labeled in the P boundary segment, and that subset is independent of the longitudinal position of the boundary. (C) Three segments in an embryo in which a single primary o blast cell was injected, showing its progeny, as inferred from a comparison of the partially labeled boundary segment to more posterior, fully labeled segments obtained when the P teloblast was labeled (as in panel A). (D) Distribution of the clone of cells labeled after injecting tracer into a single primary p blast cell, inferred from boundary segments as illustrated in panel B. Abbreviations: adc, anterodorsal cluster of central neurons; mpg, medial packet glia; pvc, posteroventral cluster of central neurons: nt, nephridial tubule; cf, cell floret. Other abbreviations as in Fig. 9. (From Weisblat and Shankland, 1985.)

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GL'NTHER S. STENT ~t a1

cell bandlets that had been labeled by injecting lineage tracer into one of the parent teloblasts (Zackson, 1982, 1984; Shankland and Weisblat, 1984; Shankland, 1987a,b; Bissen and Weisblat, 1989; Keleher and Stent, 1990). These studies revealed that each of the seven founder blast cells divides in a sequence that is idiosyncratic and stereotyped with respect to timing, orientation, asymmetry of cell division, and cell cycle composition. Furthermore, insofar as has been determined, each blast cell line has its own pattern of gene expression (Wedeen and Weisblat, 1991). Second, the fate ofsecond- and third-order m, 0, and p blast cells was ascertained by injecting lineage tracer directly into them (Shankland, l987a.b; L. Gleizer and G. Keleher, personal communications). Each higher-order blast cell was found to generate a stereotyped subset of the hemisegmental kinship group issuing from its parental primary blast cell. In the majority of cases, each subset still contributed to more than one tissue; e.g., higher-order o and p blast cells produce both neurons and epidermal cells, whereas m-derived blast cells produce both muscle fibers and nonmuscle cells. Division of higher-order blast cells d o not, therefore, lead to the typological segregation of cell fates in any simple manner. In this way, these divisions resemble the earlier blastomere divisions that generate the four ectodermal precursor teloblasts. One might conclude from the results just described that the mesodermal and ectodermal tissues of each hemisegment arise as clones founded by seven primary blast cells. In fact, the situation is more complicated, as was first inferred from results of label boundary experiments (Weisblat and Shankland, 1985) and then was demonstrated directly by injecting lineage tracer into individual primary blast cells (Shankland, 1987a,b; L. Gleizer and D. K. Stuart, personal communications). Each m, nt, o, p, or qr primary blast cell contributes a stereotyped subset of its progeny to each of several successive morphologically defined hemisegments. Progeny o f each nr, 0, p, or q, blast cell are spread over two successive hemisegments, whereas progeny of each m blast cell are in at least three successive hemisegments. Thus, each morphologically defined hemisegmerit contains the progeny of not one, but of two or three primary blast cell clones, which interdigitate across segmental borders. By contrast, the contributions of the n, and q, primary blast cells appear to be confined to single hemisegments and do not cross segmental borders.

C. ORIGIN OF

THE

SUPRAESOPHAGEAL GANGLION

Thie supraesophageal ganglion is the only component of the leech CNS that fails to be labeled after tracer is injected into any of the five

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teloblasts (Weisblat et al., 1980b). Hence, the anteriormost cells of the CNS must arise from a source other than the blast cells of the germinal bands. T h e most likely alternative source for this tissue appeared to be some or all of the 25 micromeres that arise during early cleavage: 3 each from the A, B, and C quadrants and 16 from the D quadrant (Sandig and Dohle, 1988; Bissen and Weisblat, 1989). Lineage tracer experiments confirmed this hypothesis and showed that micromeres a‘, b‘, c’, and d’ contribute their progeny to the supraesophageal ganglion (Weisblat et al., 1984). More specifically, the c’ and d’ micromeres generate bilaterally symmetric sets of supraesophageal neurons, as well as some nonneuronal cells that include the provisional epithelium of the stage 8 embryo, the prostomial epidermis, and the muscle cells of the proboscis (F. Ramirez, personal communication). Another set of micromeres, designated nopq” (Fig. 8) appears to contribute neurons in the anterior region of the otherwise teloblast-derived subesophageal ganglion (S. Ranganathan, D. K. Stuart, and L. Gleizer, personal communication). T h e observation that the supraesophageal ganglion and circumesophageal connectives of leeches are derived from micromeres, rather than from blast cells in the germinal bands, carries phyletic significance. In another major class of annelids, the polychaetes, the highly developed and complex supraesophageal ganglion is obviously a prostomial organ. It not only lies in a region that is clearly rostra1 to the metameric body segments, but it also arises developmentally as the neural tissue of a nonsegmented larva (itself derived from micromeres) to which the teloblast-derived, segmental ventral nerve cord is added later, during metamorphosis (Dawydoff, 1959). By contrast, the status of the much less elaborate supraesophageal ganglion of leeches, whose development lacks a free-living larval stage complete with its own nervous system, has long been the subject of controversy. Whitman (1887) originally conjectured that the leech’s supraesophageal ganglion is derived from the micromeres rather than from the germinal bands. Modern lineage tracers have confirmed this hypothesis, although Whitman abandoned it when he concluded that the anteriormost ganglion is a homologue of the segmental ganglia rather than a prostomial organ (Whitman, 1892). D. TRANSFATING The analyses of cell lineages described thus far indicate that leech neurogenesis is, on the whole, highly “determinate,”in the sense that the developmental fate of a particular embryonic cell in a normal embryo

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can be reliably predicted. One exception to this general finding is the indeterminacy of the fates of the O/P sister teloblasts in Helobdella. These teloblasts arise from an apparently symmetric division of their mother cell, OP, and they are morphologically as well as genealogically indistinguishable. One of the O/P sisters, designated the “generative 0 teloblast,” gives rise to the 0 kinship group, whereas the other, designated the “generative P teloblast,” gives rise to the P kinship group (Figs. 9 and 11). Whether a particular O / P teloblast takes on the role of the generative 0 o r generative P tebbidst depends on the relative position of its bandlet of daughter blast cells within the germinal band (Weisblat and Blair, 1984). T h e O/P sister teloblasts thus form an “equivalence group” (Sulston and White, 1980) of pluripotent embryonic cells. T h e following rule governs the fate taken on by the sister teloblasts: whichever O/P bandlet lies next to the q bandlet when it enters the germinal band will give rise to the elements of the P kinship group, whereas the O/P bandlet lying next to the n bandlet will give rise to the elements of the 0 kinship group. Once its blast cell bandlet is in place, the fate of either OIP teloblast is determined, in that its fate as 0 or P can be predicted reliably based on the relative position of its bandlet in the germinal band. Although under normal conditions the fate of an O/P teloblast and of its blast cell bandlet becomes determined when the bandlet enters the germinal band, the primary blast cells have not yet lost their ability to switch fates at that time (Weisblatand Blair, 1984). This fact came to light when an 0 / P teloblast, identified as the generative P teloblast by the position of its bandlet containing the first dozen or so blast cells at the time, was ablated by intracellular injection of DNase, aborting further production of blast cells. At stage 10, in the posterior regions of the embryo that had been deprived of one of the p bandlets, the ipsiiateral primary o blast cells had changed their developmental fate and had given rise to cells of the P kinship group instead (Fig. 12). Such an experimentally induced change in developmental fate-of 0 bandlet cells to a P fate-has been designated “transfating.” By contrast, an equivalent ablation of the generative 0 teloblast was found to have little effect on the fate of the primary p blast cells, which continue to follow their normal P fate. These findings have been interpreted in several ways: that (1) the blast cell descendants of an O/P teloblast are developmentally pluripotent, i.e., capable of giving rise to either 0 or P fates; (2) these blast cells will take on one o r the other fate depending upon whether they lie next to the n o r q bandlet within the germinal band; and (3) the t w o fates form a developmental hierarchy, because if there is only one O/P

LEECH NERVOUS SYSTEM DEVELOPMENT NERVE CORD SEGMENTS

xv

-

149

BODY WALL

....

....””

’...., .....:

xv I XVll XVlll XIX

xx XXI FIG. 12. 0-to-P transfating. Camera lucida tracing of seven segments of a stage 10 embryo of Helobdella triserialir. Lineage tracer was injected into the generative 0 teloblast at stage 7; some hours later, after a number of labeled o blast cells had been produced, the generative P teloblast was killed. Labeled epidermal cells are stippled; other labeled cells are black. The nerve cord is drawn separately; its true position relative to the body wall is indicated by a dotted outline. The upper segments (XV-XVII) are populated by a full complement of cells, including progeny of p blast cells, that were born before the P teloblast was killed. Here, the labeled progeny of the 0 teloblast form a typical 0 pattern. Because of the ablation, the lower segments (XIX-XXI) lack progeny of the P teloblast. Here, progeny of the 0 teloblast form a typical P pattern; np, nephridiopore complex; other abbreviations as in Fig. 9. The segment labeled XV is M11 by the enumeration scheme used elsewhere in this paper. (From Shankland and Weisblat, 1984.)

teloblast-derived bandlet present, that bandlet will assume the P fate (said to be the “primary” fate of the equivalence group) (Weisblat and Blair, 1984). Hence, in a normal embryo, the o bandlet is diverted from the primary P fate to the “secondary”0 fate by some form of interaction with the p bandlet that lies beside it. Interactions with the n and q bandlets appear to have no role in this transfating; despite ablation of the N and/or Q teloblasts, the o bandlet cells still take on the 0 fate as long as the p bandlet is present, and they are transfated to the P fate if

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the p bandlet is ablated (Weisblat and Blair, 1984; Zackson, 1984). On the other hand, the epithelial cells overlying the germinal bands may play a role in the fate-determining interactions of the o and p bandlets, because producing localized photodynamic lesions in that epithelium at stage 7 produces partial transfating of o blast cells to the P fate, even in the presence of the adjacent p bandlet (Ho and Weisblat, 1987). Transfating experiments in Theromyzon have provided additional insights into the O-to-P transfating process. Unlike in Helobdella, when the generative P teloblast is ablated in Theromyzon, transfating of the o bandlet occurs in some, but not all, segments deprived of the p bandlet (Keleher and Stent, 1990). When these embryos were observed at stage 7 or 8, in some segments that lacked the p bandlet the o bandlet had moved over into the position of the absent p bandlet, and in others it had not moved. Scoring the same embryos at stage 10 for the development of 0 or Y fates indicated that transfating had occurred only in the segments in which the o bandlet had moved into an abnormal position next to the q bandlet. There was no transfating in segments in which the o bandlet remained in its normal position next to the n bandlet. These results suggest that positional cues derived from cells other than those belonging to the n, 0, p, o r q blast cell bandlets play a role in determining the fate of the pluripotent o and p blast cells. T h e underlying m bandlet cells may provide these signals. Using the photosensitizing lineage tracer FDX, it is possible to show that the o blast cells ultimately do become committed to the 0 fate and no longer respond to ablation of their apposed p bandlet by transfating (Shankland and Weisblat, 1984). In this experiment, the generative P teloblast was injected with FDX and the generative 0 teloblast was injected with a nonphotosensitizing tracer, such as RDX or HRP. At progressively later developmental stages, germinal bands containing these photosensitized blast cell bandlets were illuminated, ablating the p bandlet cells and their descendants. By the end of stage 10, many labeled o blast cell clones in these embryos had become committed to the 0 fate. However, this commitment occurred in a sequence of three successive steps, rather than in a single event affecting the fate of the entire descendant clone of each o bandlet cell. In each of the three steps, members of the o blast cell clone become committed only to a particular portion of the entire 0 fate, concomitantly losing their potency to transfate into a particular subset of the elements of the P fate (Shankland and Weisblat, 1984).T h e three commitment steps to the 0 fate, and the corresponding losses of pluripotency, appear to be associated with successive divisions of the o blast cell clone. In each of these divisions, one of the two daughter cells becomes committed to produce a subset of 0 fate ele-

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ments, while the other daughter cell remains pluripotent for the remainder of the, as yet uncommitted, elements of the 0 and P fate (Shankland and Stent, 1986). This orderly, stepwise sequence of partial commitments accompanying asymmetric cell divisions must reflect important aspects, as yet undetermined, about the molecular nature and organization of the determinants that commit embryonic cells to their eventual fates.

V. Myogenesis and Neurogenesis

A. MYOGENESIS In all leech species studied, the musculature of the leech body wall develops mainly during growth and expansion of the germinal plate. T h e outer layer of circular muscle fibers (Fig. 1B) and the inner layer of longitudinal fibers are first to appear; shortly thereafter, the intermediate layer of oblique muscle fibers appears between the circular and longitudinal layers. Observing the development of the musculature has been aided greatly by the isolation of a monoclonal antibody that binds selectively to leech muscle fibers (Zipser and McKay, 1981). T h e circular, longitudinal, and oblique muscle layers all arise according to the same morphogenetic scheme: scaffold cells lay down the basic architecture of the layer and myoblasts then collect around the scaffold cells to establish the definitive pattern of fascicles. The scaffold for the circular and longitudinal muscle layers of the body wall is provided by a segmentally iterated gridwork of individually identifiable founder fibers (Stuart et al., 1982; Torrence and Stuart, 1986). The first circular founder fiber in each hemisegment, the primary circularfiber (Fig. 13), appears late in stage 8 or early in stage 9 and spans the entire distance from the ventral midline to the lateral margin of the germinal plate. In accord with the rostrocaudal gradient that characterizes development of the germinal plate, this fiber is first seen in anterior segments (at late stage 8) and only later in more posterior segments (at early stage 9). Each primary circular founder fiber lies along the posterior margin of the intersegmental septum, and by convention it defines the anterior edge of a segment. T h e second circular founder fiber appears soon after the primary circular fiber. This fiber is designated the septalfiber, because it lies along the anterior margin of' the intersegmental septum. Additional circular muscle founder fibers (Fig. 13) appear progressively throughout the remainder of stage 9 and into stage 10.

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FIG. 13. Myogenesis and neuroblast migration in 14 segments of a stage 9 embryo of Theromyzon nrde. The entire width of the germinal plate is shown in this tracing from a fluorescence photomicrograph. Immunolabeled muscle fibers are stippled; lineage-tracerlabeled cells of the q bandlet (on one side only) are solid black. The longitudinal extent of one segment is indicated by the bracket labeled s, at the left. The increasing numbers of circular (horizontal) and longitudinal (vertical) muscle fibers, from the posterior segments at the bottom toward the older, more anterior segments at the top, reflect the orderly appearance of new fibers over developmental time. In the posteriormost segments, the labeled q bandlet is still coherent. In more anterior segments, major (M) and minor (m) groups of labeled cells leave the q bandlet and migrate into the future ganglia (go, dashed outline), which are outlined by the primary circular (pcf) and deep longitudinal (dlf) muscle fibers; scf, septa1 circular muscle fiber. Scale bar = 50 pm. (Drawing by S. A. Torrence.)

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Longitudinal muscle founder fibers begin appearing along the lateral margins of the germinal plate very soon after the consegmental primary circular fiber (Fig. 13). At about the time that the septa1 circular fiber first appears, a prominent band of longitudinal founder fibers also appears that lies deeper in the germinal plate (i.e., further from the epidermis) than do other longitudinal fibers of the body wall musculature. These are referred to as the deep longitudinalfibers, although by stage 11 they come to lie at the same depth as the rest of the longitudinal fibers. Additional longitudinal muscle founder fibers appear progressively throughout the remainder of stage 9 and into stage 10. The architecture of the oblique muscle layer of the hemisegment is provided by a single scaffold cell, the C-cell (Jellies and Kristan, 1988a; Jellies, 1990). In Hirudo, this cell grows about 70 parallel processes, which lie at an angle of 45" to the long axis of the embryo (Fig. 14A). These processes grow out between the layers of longitudinal and circular muscle precursors and they serve as the scaffold on which myoblasts gather and orient themselves to form individual oblique muscles (Fig. 14B).The spacing between the processes appears to depend on interac-

C-cells

FIG. 14. Morphology of the C-cells, precursors of the oblique muscles, in Hirudo medicinal&. (A) Tracings of HRP-filled C-cells in one segment of E12. The ventral midline of the germinal plate runs down the ganglion (indicated by dashed lines), and the nephridiopores are about two-thirds of the way to the lateral edge of the germinal plate. (Tracings provided by D. .M. Kopp.) (B) Diagram showing how myocytes line up along the processes of C-cells to form the oblique muscles in two midbody segments at about E13. (Drawing provided by J. Jellies.)

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tions among growth cones at the ends of the processes. T h e processes of C-cells in adjacent segments are transiently linked by gap junctions at the intersegmental border. Interactions via these junctions appear to establish the extent and spacing of the oblique muscles in successive segments, thus producing a uniform distribution of oblique muscles along the whole length of the body. Myoblasts begin aggregating around their scaffolds during stage 11 of glossiphoniid development. The myoblasts then elongate and differentiate into definitive muscle fibers. The orthogonal grid of founder fibers thus serves as a template for the development of the definitive muscle fascicles of the adult. It is not known whether the founder fibers for the circular and longitudinal muscles persist into the adult, but it appears that the C-cells die after completing their task of organizing the oblique muscle layer (Jellies and Kristan, 1988a). The use of scaffold cells for establishing muscle layers is a developmental strategy also encountered in the development of insects (Ho et d., 1983) and crustaceans Uellies, 1990).

B. GANGLIOGENESIS

1. Ganghonu: Rluliments The process of ganglion formation is similar in hirudinid (Fernandez and Stent, 1983) and glossiphoniid (Fernandez, 1980; Kuwada and Kramer, 1983; Torrence and Stuart, 1986; Stewart et al., 1987) leeches. The following detailed description is based primarily on observations of glossiphoniid embryos. In stage 8 and early stage 9, when the germinal plate of a glossiphoniid embryo has just been formed by the coalescence of the right and left germinal bands, its constituent bandlets are still coherent columns of cells. T h e four bilateral pairs of ectodermal bandlets lie in a single layer, and the single bilateral pair of mesodermal bandlets lies just below the ectodermal bandlets (Fig. 6). At this stage, before there are morphologically recognizable ganglionic rudiments, the primary circular and deep longitudinal muscle fibers described in the previous section first become detectable. When ganglionic rudiments appear slightly later in stage 9, each occupies the rectangular territory bounded by two successive primary circular fibers and by the right and left deep longitudinal fibers (Fig. 13). Thus, these muscle fibers outline the presumptive ganglionic territory in each segment. At the beginning of gangliogenesis, cells begin accumulating within

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the presumptive ganglionic territory by proliferation of cells locally and by migration from more lateral parts of the germinal plate. In the lateral regions of the presumptive ganglionic territories, bilaterally paired aggregations of cells form; they are the earliest visible morphological sign of ganglion formation (Fig. 7A). In accord with the rostrocaudal developmental gradient in the germinal plate, these cell aggregations appear first in the anteriormost segments and progressively later in ever more posterior segments. The longitudinal muscle fibers that will extend along the interganglionic connective nerves appear at about this stage as flat cells, stainable with the antimuscle antibody, on the deep (future dorsal) aspect of the nascent ganglia. Where it overlies the primary circular muscle fibers, the presumptive neural tissue is thinner, forming narrow clefts that demarcate the margins of individual ganglia. Adjacent ganglia remain in contact with one another throughout most of stage 9, during which time longitudinal nerve fiber tracts are established between them. Thus, when the midbody ganglia begin to move apart from one another late in stage 9, they are already linked via connective nerves. As the embryo elongates during later stages, the midbody ganglia move increasingly far apart and the connective nerves lengthen accordingly. Rather than separating, the anteriormost four and posteriormost seven ganglia fuse to form the subesophageal and caudal ganglia (Fernandez, 1980; Kuwada and Kramer, 1983; Torrence and Stuart, 1986). Ganglionic neurons are parceled into their six glial packets in stage 10 (Kuwada and Kramer, 1983). As in the vertebrate nervous system (Oppenheim, 1991), neuronal cell death helps to shape the developing nervous system. Studies of neuronal cell number during development of the hirudinid Huemopis mumnoruta (Stewart et ul., 1986) have shown that by E10, newly formed midbody ganglia contain approximately 14% more neuronal cell bodies than does an adult ganglion. Between El0 and E20, this number declines to the adult value. The presence of pycnotic and fragmenting cells during this decline indicates that cell death is probably responsible for the decrease in cell number. Cell death may perform three functions in nervous system development (Truman, 1984): ( 1 ) to regulate the size of neuronal populations, serving to match cell numbers between two regions of the CNS o r between the CNS and the periphery; (2) to remove unneeded cells whose production is a by-product of fixed cell lineages; and (3) to remove cells whose function is transient, after they are no longer needed. It is likely that cell death plays all of these roles in the development of the leech nervous system (Stewart et al., 1986), as illustrated by the following examples. First, cell death regulates cell number by removing some neu-

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rons that arise as pairs but are unpaired in adult ganglia. Second, an immunochemically identified pair of putative neurons that arise in each segmental ganglion of Hirudo appear to be transiently functional. These cells grow processes, persist for about 2 days, and then die (Stewart et al., 1987). Third, it seems likely that the stereotyped cell lineages of the leech generate the same complement of neural precursor cells in each segment, and that segment-specific patterns of cell death play some role in producing distinct segmental identities among midbody ganglia. The genesis of extra neurons also contributes to unique, segmentspecific characteristics of some ganglia. As mentioned above, ganglia in the reproductive segments, M 5 and M6, of adult leeches contain significantly more neurons than d o other midbody ganglia (Macagno, 1980). The origin of the increased cell number in ganglia of reproductive segments M5 and M 6 has been studied in embryos of the hirudinid Hnemopt-s. Here the ganglia in reproductive segments contain the same number of neurons as other midbody ganglia until E20 (Stewart et al., 1986). Thereafter, neuronal cell number gradually increases in ganglia of segments M 5 and M 6 through late embryonic and postembryonic development, while the cell number remains constant in other midbody ganglia. Immunohistochemical studies of ganglionic DNA replication indicate that the additional cells are generated by proliferation of neuroblasts within ganglia of the reproductive segments (Baptista et al., 1990). 2 . Neuroblmt Migration Migrations of neuroblasts and immature neurons have long been recognized as playing crucial roles in the development of vertebrate nervous systems (Rakic, 1972; Le Douarin, 1984), but they have been much less prominent in accounts of invertebrate neurogenesis. Attention was first drawn to the importance of cell migration in the neural development of glossiphoniid leeches when it was observed that all blast cell bandlets contribute cells to the CNS, although the p and q bandlets initially lie completely outside the presumptive ganglionic territory. Hence the progeny of p and q bandlet cells that are destined for the CNS must migrate to enter the ganglionic rudiments (Weisblat et a/., 1984). Studies analyzing the migrations of lineage-tracer-labeled cells have revealed that in each segment, small groups of cells follow stereotyped migration pathways from the site of their birth to their characteristic final positions. Cells of the q bandlet must migrate farther to reach the CNS than do any other ectodermal neural precursor cells and their migrations have been described in the greatest detail (Weisblat et al., 1984; Torrence and

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Stuart, 1986; Torrence, 1991) (Fig. 13). The description of the fates of these migrating cells reemphasizes the diversity of cell fates that are assumed by the progeny of the ectodermal bandlet cells. In every hemisegment, two small groups of cells leave the q bandlet to migrate toward the presumptive CNS. These cells divide several times during and after their migration. T h e major group migrates toward the ventral midline along the anterior margin of its segment, just posterior to the primary circular muscle fiber. Most of the cells of this group enter the ganglionic rudiment, where they give rise to particular glia and neurons (cf. Fig. 9). A few cells remain outside the ganglionic rudiment and they contribute to a cluster of peripheral neurons in the segmental MA nerve. The remainder of this peripheral cluster is derived from the smaller, and later-departing, minor group of migratory cells that also divide during their migration. These cells migrate medially along a midsegmental path, posterior to the path followed by the major group. After the peripheral cluster has formed, one cell leaves it to enter the ganglion. The q bandlet cells that d o not migrate give rise to the dorsal epidermis and peripheral neurons. A single group of cells in each segment migrates from the p bandlet toward the presumptive CNS, starting out at about the same time as the migratory q bandlet cells. Like the minor group of q bandlet cells, migratory cells from the p bandlet follow a midsegmental path, approximately halfway between successive primary circular muscle fibers. These cells will form cell floret 1 in the periphery, as well as central neurons and a packet glial cell (cf. Fig. 10) (Torrence and Stuart, 1986; Braun and Stent, 1989a). Most cells of the n and o bandlets arise within the presumptive ganglionic territory and are incorporated directly into the ganglionic rudiments. Nevertheless, they also undergo characteristic rearrangements and short migrations within the ganglionic rudiment to reach their definitive positions (Stuart et al., 1987; Braun and Stent, 1989a). For example, in the adult midbody ganglion, the serotonin neuron pms is positioned at the posterior margin of the segmental ganglion, although it arises from a primary blast cell clone that initially populates the anterior ganglionic quadrant (Stuart et al., 1987; Bissen and Weisblat, 1987). Thus, pms o r its precursor must migrate posteriorly a distance of at least one-half segment to reach its definitive position. A few central neurons turn out to be of mesodermal provenance (Fig. 10) (Weisblat et al., 1984; Kramer and Weisblat, 1985). They arise as a group of five morphologically immature, m bandlet progeny that migrate into each nascent hemiganglion to take up their specific positions (Shankland and Martindale, 1989). Some prospective peripheral neu-

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rons also must migrate to reach their characteristic positions. For example, peripheral neurons nzl, nz2, and nz3 (collectively referred to as nz neurons) (Fig. 9) arise within the presumptive ganglionic territory and migrate laterally to reach the periphery. Furthermore, several o or p bandlet-derived peripheral neurons (pz6, pzl0, and LD2) arise near the lateral margin of the presumptive ganglionic territory and migrate laterally to reach their definitive positions (Weisblat et al., 1978; Braun and Stent, 1989a). Precursors of nonneural ectodermal cells have also been shown to undergo similar migrations (Martindale and Shankland, 1988; Braun and Stent, 1989a). In summary, the determinate patterns of cell division that generate the neurons and glial cells of the leech nervous system do not place most cells directly into their characteristic final positions. Instead, the definitive pattern of neuronal positions is produced by extensive and stereotyped cell rearrangements and migrations. 3 . Tissue Interactions

To investigate the role of tissue interactions in controlling the migrations that generate the neuronal pattern of the leech nervous system, the development of lineage-tracer-labeled cells that are descended from the n and q bandlets has been studied in embryos selectively deprived of other bandlets. T h e results showed that many neurons differentiate even when deprived of their normal intractions. For example, identified neurons derived from the n bandlet, including the peripheral nz neurons and the central serotonin neurons (Figs. 3 and 10; also, see later, Fig. 19), found their characteristic positions, grew axons, and (in the case of the serotonin neurons) neurochemically differentiated in segments deprived of the o and p bandlets or of the q bandlet (Blair and Weisblat, 1982; Blair, 1983; Stuart et al., 1987, 1989). Similarly, cells derived from the q bandlet followed their normal migration pathways, found their normal final positions, and (in the case of the peripheral dopamine neuron MD) neurochemically differentiated in segments deprived of n, 0, or p bandlets (Blair, 1983; Torrence, 1991). Thus, interactions among cells of different ectodermal lineages are not required for the differentiation of at least some aspects of neuronal phenotype. In contrast, two types of teloblast ablations did have dramatic effects on n bandlet progeny. Although the mirror-symmetrical patterns formed by the left and right n bandlets are normally restricted to their respective sides of the embryos (Figs. 10 and 15A) (Weisblat et al., 1984; Kramer and Weisblat, 1985; Weisblat and Shankland, 1985; Torrence and Stuart, 1986; Stuart et al., 1989),elimination of either one n bandlet (Fig. 15B, left panel) (Blair and Weisblat, 1982; Blair, 1983; Stuart et al.,

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1987, 1989) o r one m bandlet (Fig. 15C, left panel) (Blair, 1982; Torrence et al., 1989) induces an abnormal cross-over of n bandlet cells to the opposite side. After n bandlet ablation, one hemiganglionic complement of several identified cell types was missing, including neuropil glial cells (Blair and Weisblat, 1982), serotonin neurons (Blair, 1983; Stuart et al., 1987, 1989), and the peripheral neurons nzl, nz2, and nz3 (Stuart et al., 1989), showing that descendents of the absent n bandlet were not regulatively restored from any other source. Furthermore, the single surviving n bandlet generated its normal segmental complement of descendant cells, but they were abnormally distributed, with some cells on one side of the ganglionic midline and some cells on the other. When peripheral nz neurons and central serotonin neurons abnormally crossed the ventral midline into hemisegments that were deprived of an n bandlet, they nearly always occupied the normal positions of their missing homologues (Fig. 15B, right panel). This observation might suggest that pluripotent n bandlet cells differentiate into a particular neuron because they occupy a particular location. However, this hypothesis predicts at least some bilateral duplications of identified neurons when the descendants of a single n bandlet are spread over both sides of the nervous system, because two pluripotent cells should sometimes happen to occupy homologous positions on the opposite sides of the same segment. Such duplications were never observed. Therefore, cells that cross over into contralateral territory must be committed to finding the characteristic position of absent homologous neurons. [Neural precursor cells differ in this regard from epidermal precursors, which are not committed to any particular position within the epithelium (Blair and Weisblat, 1984).] T h e results described so far do not exclude the possibility that a neuronal precursor cell initially is instructed only about its destination following migration, and it receives further instructions regarding the remainder of its phenotype once it has completed its migration. This possibility has been eliminated, however, by the behavior of n bandletderived neuronal precursor cells that have been deprived of their underlying m bandlet (Fig. 15C, left panel) (Blair, 1982; Torrence et al., 1989). Stage 10 or 11 embryos that were prevented from forming most of one m bandlet contained a zone of segments lacking mesodermal tissues on one side (Fig. 15C, right panel). Just as in the case of hemilateral n bandlet ablations, when one m bandlet is missing, some precursors cross the midline and the resulting immigrant neurons occupy their characteristic sites and develop their characteristic phenotypes. In this case, however, on the side containing the m bandlet there are duplicate n bandlet-derived neurons, one native and the other immigrant. In these

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embryos, some of the n bandlet-derived cells remain on the mesodermdeprived side, where they are scattered along the nerve cord and are not organized into recognizable patterns. Nevertheless, many of the stay-athome neurons differentiated as serotonin neurons (Fig. 15C, right panel) (Torrence et al., 1989). Several general conclusions can be drawn from these experiments. First, a precursor cell does not need to occupy its normal position nor does it need contact with a normal complement of mesoderm, to differentiate neurochemically. Second, the fact that migration of neuronal precursor cells from either side is normal on the side with mesoderm and disorganized on the side lacking mesoderm implies that the posiFIG. 15. Summary of experiments analyzing the effects of unilateral N (panel B) or M (panel C ) ablation on the development of identified neurons. In each panel, the protocol is diagrammed on the left and the results are illustrated on the right in four segments from a stage 11 embryo. Round black dots within the diagrammatic ganglia represent immunoreactive serotonin neurons (cf. Fig. 3); black ovals next to the ganglia represent peripheral nz neurons (cf. Fig. 9). Both cell types are derived from the n bandlets. Hatching and stippling represent differently colored lineage tracers. Anterior is up. (A) Normal fate mapping. Protocol: the left and right n bandlets were labeled with differently colored lineage tracers [red-fluorescing rhodamine dextran amine (RDA) and the yellow-fluorescing fluorescein dextran amine (FDA)]by injecting them into their parental teloblasts (circlesat the bottom of each bandlet) at stage 6 or 7. Result: cells descended from either n bandlet were restricted to their respective sides of the nerve cord. (B) Effects of unilateral n bandlet deprivation. Protocol: the left and right n bandlets were labeled as in A. Later, after a few labeled n blast cells had been formed, further formation of the right n bandlet was aborted by killing the right N teloblast by injecting it with DNase. Result: anterior control segments (top ganglion) contained progeny of both n bandlets, born before the teloblast was killed, and were completely normal. More posterior segments (bottom three ganglia) lacked progeny of the right n bandlet, and all serotonin and nz neurons were derived from the left n bandlet. In such segments, the left n bandlet gave rise to its normal complement of identified neurons. These neurons were abnormally distributed across both sides of the nervous system, but on either side they occupied positions appropriate for their cell types. ( C ) Effects of unilateral m bandlet deprivation. Protocol: the left n and m bandlets were labeled with differently colored lineage tracers, as in A. Later, after a few labeled blast cells had been formed, further formation of the left m bandlet was aborted by killing the left M teloblast with DNase. Result: anterior control segments (top ganglion) contained progeny of a complete Complement of blast cells, born before the teloblast was killed, and were completely normal. More posterior segments lacked progeny of the left m bandlet. Here, both n bandlets gave rise to serotonin and nz neurons. Some progeny of the mesodermdeprived left n bandlet crossed into nondeprived right hemiganglia, where they intermingled with their normally contralateral homologues and found appropriate, cell typespecific positions adjacent to those homologues (middle two ganglia). Other progeny of the left n bandlet remained in the mesoderm-deprived region. Such cells were not organized into recognizable patterns. In the extreme posterior segments (bottom ganglion) progeny of both n bandlets were disorganized. (A and B from Stuart et al., 1989; C from Torrence et al., 1989.)

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tional information used by migrating neuronal precursor cells is provided by mesodermal tissue. Third, the observation that n-derived neurons can cross the ventral midline, but d o not normally do so, implies that under normal conditions the apposition of the left and right n bandlets prevents the cells of both n bandlets from crossing the midline. Fourth, because immigrant n bandlet cells from a side lacking mesoderm intermingle freely with the resident n bandlet cells, the ventral midline of a leech embryo does not represent a line of clonal restriction of the same kind as compartment boundaries in insect embryos (Garcia-Bellido et al., 1973; Crick and Lawrence, 1975). T h e migration of cells derived from the q bandlet cells, following ablation of specific tissues, suggested a further role for mesoderm in guiding cell migrations. After the mesoderm is ablated on one side of an embryo, q bandlet-derived cells begin to migrate toward the ventral midline at the normal time, but they fail to follow normal migration pathways or to find their normal definitive positions. Thus, commitment of particular q bandlet-derived cells to migrate does not depend on the presence of mesoderm, but their ability to follow normal pathways and recognize their normal destinations does (Torrence, 199 1). T h e results of these ablation experiments strongly suggest that neuronal precursor cells attend to local cues to find their normal positions, and that at least some of the cues are of mesodermal origin (Stuart et ul., 1989; Torrence et al., 1989; Torrence, 1991). Because neurons are normally arranged with bilateral symmetry in a segmentally iterated array, it is likely that the mesodermal positional cues are similarly arranged. These characteristics accord well with the developmental origin of the mesoderm from the bilaterally symmetrical m bandlets, each of which products segmentally iterated complements of progeny cells from its longitudinal series of primary blast cells. C. NEUROCHEMICAL DIFFERENTIATION

Neurochemical differentiation in leech embryos has been followed by diverse methods. To determine when neurons could first synthesize acetylcholine (ACh) or serotonin, embryonic nerve cords that had been isolated at various developmental stages were exposed to [3H]choline o r to [3H]hydroxytryptophan and their rate of ["H]ACh or [3H]serotonin synthesis was assayed. In Hirudo, detectable conversion of 3H-labeled precursors into ACh or serotonin appeared during E7-E8, when ganglia are first forming, and then rose rapidly from E9 onward (Wallace, 198 1). Similarly, Huementeriu embryos synthesize very small amounts of

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ACh and serotonin early in stage 9 (Fig. 16) (Glover et al., 1987). The amount of transmitter synthesized rose slowly in middle or late stage 9, and then rapidly duriqg stage 10. By the beginning of stage 11, transmitter synthesis was 20-fold higher than at stage 9. Staining the embryonic nervous system for the presence and distribution of acetylcholinesterase (Fitzpatrick-McElligott and Stent, 1981) indicated that the capacity to hydrolyze ACh develops in parallel with the capacity to synthesize it. Also, there is a concomitant rise in the levels of ACh and serotonin, as measured by radioimmunoassay (Fig. 16) (Glover et al., 1987). The appearance and accumulation of various transmitters in identified neurons have also been followed using specific histochemical methods. In glossiphoniids, neuronal staining with an antiserotonin antibody is first detectable at midstage 9. The number of immunoreactive serotonin neurons in the nerve cord increases rapidly until early stage 10 and approaches that of the adult complement by early stage 11 (Fig. 16C) (Glover et al., 1987). This progressive increase in the number of immunoreactive serotonin neurons has two components. One is a progressive increase in the number of ganglia containing a given type of immunoreactive serotonin neuron; in accord with the rostrocaudal development gradient, serotonin neurons of a given type appear first in the anteriormost ganglia, and neurons of the same type are added successively in more posterior ganglia. The other component is a progressive addition of cell types within any given ganglion, with the Retzius neuron appearing first and the other serotonin neurons later. Retzius neurons are labeled by an antiserotonin antibody very early during axonogenesis, whereas the other serotonin neurons cannot be labeled until their axonogenesis is well under way (Glover and Mason, 1986; Glover et al., 1987; D. K. Stuart, unpublished observation). The characteristic blue-green fluorescence induced in dopamine neurons by glyoxylic acid treatment has been used to study the accumulation of dopamine in the peripheral neurons MD and LDl (Figs. 3 and 10). In Haementeria (Glover et al., 1987),stainable MD neurons first appear in anterior segments early in stage 10, at which time they have already projected axons into the CNS. During stage 10 and early stage 11, central arbors of MD neurons become more elaborate, and additional MD neurons appear in progressively more posterior segments. LD 1 neurons cannot be labeled before stage 11, In Hirudo, MD neurons become stainable in anterior segments by about E9-El0, shortly after completion of gangliogenesis. As in Haementm.u, the MD axons in Hirudo project into the CNS by the time they contain enough dopamine to be labeled (Dietzel and Gottmann, 1988). Interestingly, in ganglia of H i d o embryos that have been preincubated in dopamine between E7 and El 1,

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FIG. 16. Development of serotonin metabolism in Haementrrin gliilianii during stages 911. Each interval on the ordinate corresponds to 1 day of-development, with the first time point being taken early in stage 9 (approximately 10 days after egg deposition for this species) and the last early in stage 11. (A) Serotonin content of nerve cord (circles) and body wall (squares), i.e., germinal plate from which the nerve cord had been dissected. The asterisk represents the value obtained for a total germinal plate, from which the nerve cord was not dissected, at the earliest time point, showing that serotonin was undetectable at that stage. Error bars indicate standard deviation (SD) of the mean; if the SD was smaller than the height of the symbol. no error bar is shown. (B) Specific serotonin content, in picomoles of serotonin per microgram of protein; same data as in A. (C) The number of serotonin-irnmunoreactive neurons in the nerve cord. The dashed horizontal line indicates the adult complement of serotonin-containing neurons. (D) Total capacity for synthesis and accumulation of serotonin by the nerve cord, as measured in an electrophoretic assay following a 3-hr incubation in tritiated serotonin. (E) Specific capacity for synthesis and accuniulation of serotonin by the nerve cord per microgram of protein. (From Glover et al., 1987.)

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serotonin neurons have the blue-green fluorescence characteristic of dopamine following glyoxylic acid treatment (Dietzel and Gottmann, 1988). After E l 1, serotonin neurons gradually cease to stain for dopamine after dopamine preincubation. Thus, it appears that the developing serotonin neurons first acquire, and then lose, the ability to take up dopamine. GABAergic neurons, including the inhibitory motor neurons innervating the longitudinal muscles (Cline, 1983, 1986), possess high-affinity GABA uptake systems that provide rapid resorption of the transmitter from the synaptic cleft (Schousboe, 1981). The development of this uptake system in Huementeria has been studied by autoradiography of embryonic ganglia from various stages after incubation of the nerve cord in [3H]GABA. A pair of cell bodies that takes up [3H]GABA first appears in anterior ganglia at early stage 9. In a pattern similar to serotonin neurons, at progressively later times homologous GABAergic cells appear in increasingly more posterior ganglia, while within a given segmental ganglion, additional GABAergic cells appear in a stereotyped order until early in stage 11. By that time, the full adult complement of about 30 neurons that take up 13H]GABA is present in each ganglion (Glover et ul., 1987). Another set of adult neurons is labeled by an antibody directed against the molluscan small cardiac peptide B (Shankland and Martindale, 1989; Evans and Calabrese, 1989). In the following discussion, these neurons will be referred to as “SCP neurons,” although the leech neuropeptide that is recognized by this antibody is probably not identical with molluscan SCP (Evans and Calabrese, 1989). Many SCP neurons are also labeled by an antibody against the neuropeptide FMRFamide. In Theromyzon and Helobdellu, the neurons labeled by anti-SCP or antiFMRFamide antibody first appear late in stage 10 in anterior segments and near the beginning of stage 11 in posterior segments. They appear, therefore, considerably later than most serotonin neurons. The axons of SCP neurons in the interganglionic connective become immunoreactive at the same time as their cell bodies, suggesting that, unlike Retzius neurons, SCP neurons project axons before accumulating detectable amounts of neurotransmitter. Among the earliest SCP neurons to stain are the phenotypically asymmetrical rostra1 alternating SCP (RAS) (in anterior segments) and caudal alternating SCP (CAS) (in posterior segments) neurons, whose development will be discussed in more detail below. T h e number of immunoreactive cell bodies increases as development proceeds, and some cells appear that are labeled by the antiFMRFamide antibody, but not by the anti-SCP antibody (Shankland and Martindale, 1989).

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A

Adult

Late stage 9

-

Early stage 10

Early stage 11

-.

/-

C

FIG 17. Development of electrogenesis and peripheral processes in the mechanoreceptive P cells of H a m t i t e r i n ghilianii. (A) Development of electrogenesis. The adult form of the action potential is shown in the top trace. N o active electrical responses could be elicited before the last day of stage 9, when a small voltage-dependent response is present on the largest depolarization shown in the leftmost trace. Early in stage 10 (middle trace), a response resembling an action potential can be elicited, which becomes progressively faster, larger in amplitude, and with an after-potential during stage 1 I (rightmost trace). All responses were elicited by passing current into the neurons during the time between the downward deflections near the beginnings of the traces and the upward deflections near the end of the traces. The calibrations apply to all four traces. (B) Development of the‘peripheral processes of Pv from late stage 9 to early stage 11. The outlines of the ganglia are shown in stages 9 and 10,but are left out of the stage 1 1 drawing in order to show the processes of PV,which pass under the ganglion. (C) The corresponding develop-

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In summary, neurochemical differentiation begins soon after the ganglion have formed in stage 9, but proceeds mainly during the expansion of the germinal plate and elaboration of the ganglia in stage 10. Each neuronal cell type undergoes neurochemical differentiation according to a characteristic schedule-some types begin to differentiate very soon after ganglion formation and others wait for another week or so. The early appearance of some aspects of neurotransmitter metabolism in leech embryos, coinciding with-or even preceding-the earliest stages of process outgrowth (cf. Section V , D ) , has led to the suggestion that neurotransmitters may participate in the regulation of neuronal or even of general development, in addition to their subsequent role in synaptic transmission (Fitzpatrick-McElligottand Stent, 1981; Glover et al., 1987). Such regulation has, in fact, been suggested in other organisms (e.g, Buznikov et al., 1968; Kusano et al., 1977; Lauder and Krebs, 1978; Haydon et al., 1984).

D.

ELECTROPHYSIOLOGICAL DIFFERENTIATION

To follow the electrophysiological differentiation of glossiphoniid leech neurons, intracellular recordings were made from embryonic neurons, glial cells, and muscle fibers in a germinal plate preparation viewed with transillumination under differential interference contrast (Nomarski) or fluorescence optics (Kuwada and Kramer, 1983; Kramer and Stent, 1985). By early stage 10,junction potentials can be recorded from the longitudinal muscle fibers, indicating that motor neurons have innervated their peripheral targets. The development of electrogenic properties has been followed for the mechanosensory P neurons (Kuwada and Kramer, 1983) (Fig. 17A).At midstage 9, the embryonic P neurons are electrophysiologically passive and their resting potentials and input resistances are -40 to -60 mV and 200 to 500 MR, respectively, as compared to values of -35 to -45 mV and 100 MR in adult neurons. By late stage 9, the mechanosensory P neurons begin to respond actively to passage of depolarizing current into the cell body, first with delayed rectification and then with an all-or-none depolarizing ment of PD. The images in B and C are tracings made from representative Lucifer Yellow fills of P cells; the size calibration applies to all the tracings. The dorsal (DM), lateral (LM), and ventral (VM) midlines are indicated in the drawings of the cells at stages 10 and 1 1 . The size calibration applies to all drawings in B and C. (A and B from Kuwada and Kramer, 1983; C based on data from Kuwada and Kramer, 1983, and Kramer and Kuwada, 1983.)

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transient. Early in stage 11, by which time process outgrowth is well under way, P neurons respond to depolarization with an overshooting action potential that resembles the characteristic action potential of the adult cells. However, the action potential lasts much longer in the embryonic than in the adult cells and can be extended even further by bathing the preparation in high-concentration Ca2+ saline. T h e P neuron action potential takes on the fully adult properties (Nicholls and Baylor, 1968) only during postembryonic growth of the juvenile leech. Spontaneous postsynaptic potentials, excitatory as well as inhibitory, can be recorded in the P cells from early stage 10, after their neuropilar processes have started to develop. Thus, formation of functional synapses in the CNS begins at about the same time as formation of neuromuscular junctions in the periphery. A similar time course of electrophysiological development has been found also for the excitatory L motor neuron of the longitudinal muscles (Kuwada, 1984). E. MORPHOLOGICAL DIFFERENTIATION To follow the morphological differentiation of leech neurons, especially the pattern of axonal outgrowth, horseradish peroxidase (Muiler and McMahan, 1976) o r Lucifer Yellow dye (Stewart, 1978) has been injected intracellularly via microelectrodes at various stages to reveal the anatomy of individual embryonic neurons (Kramer et al., 1985; jellies and Kristan, 1988a,b).In this way, it has been found that many cell bodies in the nascent segmental ganglia of glossiphoniids have not yet grown any processes late in stage 8. However, as judged by the intercellular passage of injected dye (Stewart, 1981), at this stage sets of cell bodies are dye coupled, implying that they are connected by gap junctions. By early stage 9, exuberant growth of fine processes is under way, and the dye coupling has largely disappeared. [Early dye coupling of cell bodies, followed later by its loss, occurs also among embryonic grasshopper neurons (Goodman and Spitzer, 1979).] T h e initial fine processes disappear by midstage 9, having been replaced by a single process that, for most cells, is directed toward the midline of the ganglion, traversing the region of the future neuropil. By late stage 9, the processes of particular cells have established the trajectories of the future segmental nerves and the connectives (Kuwada, 1982; Kuwada and Kramer, 1983). 1. Mechnosensory Neurons

The characteristic projections and receptive fields of mechanosensory neurons could arise during development in either of two ways. On the

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one hand, each neuron might first project axonal processes indiscriminately into all four major segmental nerves-AA, MA, DP, and PP-followed by later pruning of inappropriate processes through retraction or degeneration. On the other hand, each neuron might, from the very beginning, project only appropriate axonal processes (i.e., those that are retained into adulthood). The first possibility implies that specific axonal projection patterns arise by selective survival of haphazard axonal outgrowth, a process that might be mediated by interaction between axons and their appropriate targets. The second possibility suggests that specificity arises by directed outgrowth toward targets, a process mediated by recognition of landmarks on the growth substratum. In order to elucidate which of these mechanisms shapes the receptive fields of the mechanosensory neurons, the axonal arborization patterns of neurons P, and P, were examined at progressively later embryological stages (Fig. 17B and C) (Kuwada, 1982; Kuwada and Kramer, 1983; Kramer and Kuwada, 1983; Kramer and Stent, 1985). Like other leech neurons, these two neurons arise as morphologically undifferentiated neuroblasts in the ganglionic primordium. They can be identified in midbody ganglia of stage 9 embryos by their relatively large size and characteristic location. The first process emerges from the P, or P, cell body at its medial pole and grows toward the midline of the future ganglion neuropil. Just before reaching the midline, the process forms a T-junction, with one arm of the T growing toward the next anterior and the other arm growing toward the next posterior ganglion, via the connective nerve. In the case of cell P,, several additional, peripherally directed processes emerge concurrently from the lateral pole of the cell body and grow into the future ventral germinal plate bordering the ganglion. By late stage 9, one peripheral process of the P, neuron has grown far enough laterally to reach the future dorsal germinal plate. This process extends few branches before reaching dorsal territory, even though it is growing across uninnervated territory. Apparently the axon fails to branch because it grows along the surface of a broad, flat cell that extends from the edge of the ganglion to the future lateral edge of the embryo (Kuwada, 1982). This process will become the main adult axon of the P, neuron, which runs in the DP nerve. The other peripheral processes of the P, neuron, which have remained in the future ventral territory, disappear. Some of the ventral skin is eventually innervated by the P, neuron via an axon in the MA nerve, a process that forms as a branch of the central axon. By contrast, the P, neuron grows only one peripheral process that normally emerges from the lateral pole of the cell body and grows into the future ventral germinal plate. This process

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will become the main adult axon of the P, neuron, running in the MA nerve. The minor receptive fields of these neurons are formed as branches of the central P, and P, axons, which extend in the connectives to the adjacent ganglia and exit from nerves in these ganglia. These processes in adjacent segments form innervation fields similar to the major fields that are being formed simultaneously by the local P, and P, neurons. Meanwhile, the central processes also begin to proliferate neuropilar processes within the ganglia, to provide for the eventual network of synaptic connections. It should be noted that although both P neurons arise as bipolar neurons, with one (central) process emerging from the medial pole of the cell body and another (peripheral) process emerging from the lateral pole, they become monopolar by late stage 10. T h e conversion from bipolar to monopolar morphology of embryonic leech neurons is the result of peripheral and central axons merging at a Tjunction, where they remain attached to the cell body via the single axon that forms the stem of the T. A similar bipolar-to-monopolar conversion occurs in embryonic dorsal root ganglion cells of the vertebrate spinal cord (Tennyson, 1965). Like axons of many other embryonic neurons (Ramon y Cajal, 1929), the developing central and peripheral P cell processes carry numerous filopodia, both at their growth cone tip and on their sides. These filopodia are much more abundant on the peripheral processes that grow out first, directly from the cell bodies, than on the peripheral processes that grow out later from the central processes, either in the ganglion of origin o r in the adjacent ganglia. T h e large number of filopodia on the initial peripheral P cell process is consistent with their putative role in pioneering the peripheral pathways, because numerous filopodia are typical of growth cones at pathway choice points (Tosney and Landmesser, 1985; Caudy and Bentley, 1986). At the time the P, axons first contact skin, the germinal plate is still expanding circumferentially and large areas of the body wall have yet to form. The axon that grows peripherally from the cell body along the future MA nerve and that establishes the primary subfield appears to follow a predesignated path, because its initial peripheral branching pattern is highly stereotyped. Later, a branch of the central (neuropil) process of the P, cell may leave the ganglion via the PP nerve and establish a secondary sensory subfield. Upon circumferential expansion of the germinal plate, the peripheral axons grow and sprout additional branches to innervate new territories in the developing body wall. At no time during the outgrowth of the P, axon do any “tentative” processes enter the AA or DP nerve branches, and by the end of stage 10

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the fraction of cases in which P, processes extend into the PP branch is only slightly greater than in the adult. Hence it would appear that the adult projection pattern of this mechanosensory neuron results not from an initially haphazard projection that is selectively pruned, but rather from the specific and directed outgrowth of processes. These observations indicate, moreover, that the relationships among various components of the adult mechanosensory receptive fields are determined before the peripheral axons leave the ganglion. For example, in the major field of cell P , the primary subfield, which is the largest subfield and is invariably present, is the first to be established, and it arises by direct axonal outgrowth along the MA nerve. The secondary subfield, which is smaller and sometimes absent altogether, is established only later, by an axonal branch of neuropil processes, a branch that exits the ganglion via the PP nerve, if it forms at all. Similarly, the much smaller minor fields of cell P , within the body wall of the adjacent segments, are established by intersegmental axon branches that exit via the MA nerve of the adjacent ganglia. These branches grow out only after the primary axon that establishes the major field of the serially homologous P, neuron has already begun to innervate its own segmental body wall. This pattern suggests that some mechanism of territorial competition helps to establish the mechanosensory receptive fields, in which the earliest process to arrive has the advantage.

2. Growth Guidance by Muscle Fibers Stereotyped axonal outgrowth and mechanosensory receptive fields suggest that developing processes grow along prespecified pathways and form branches at prespecified points. The orthogonal grid of embryonic circular, longitudinal, and dorsoventral muscle fibers seems a likely source of this information, because the fibers are already in place in the germinal plate at the time axonogenesis begins. For instance, by late stage 9, some axons have formed a T-junction on reaching the midline of the ganglionic neuropil and have begun to grow longitudinally, anteriorly, and/or posteriorly out of the home ganglion and into the next ganglion. These longitudinal axons follow two pairs of the most medial longitudinal muscle fibers-founders of the future longitudinal muscle fascicles in the connective nerves. At this stage, these muscle fibers are the only overt structures connecting the ganglia (Kramer and Stuart, 1982; Stuart et al., 1982). Some interganglionic axons follow the more lateral of these muscle fibers, and thereby pioneer the paired lateral connective nerve tracts. Other axons course between the more medial muscle fiber pair and thus pioneer the unpaired connective nerve tract, or Faivre's nerve. Later, during stage 10, processes of sensory

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and effector neurons grow out of the lateral margin of the ganglion into the peripheral germinal plate, following circular muscle fibers. The axon of the P, neuron follows the septa1 circular muscle founder fiber, apparently thereby pioneering the MA segmental nerve trunk. At intersections between the circular founder fiber and particular longitudinal fibers, the outgrowing P, axon sprouts a longitudinal branch, apparently thereby pioneering the characteristic longitudinal branching pattern of the adult segmental nerve. All the higher-order axon branches of the P, neuron are also aligned with either a circular or a longitudinal muscle fiber. T h e axon of the P, neuron grows out along the most medial dorsoventral muscle precursor to the future dorsal territory of the germinal plate, among the first axons to appear in the nascent DP nerve (Kuwada, 1982).

3. Interneurons The axonal projection of the intersegmental interneuron S is of a particularly simple form, greatly facilitating observation of its development (McGlade-McCulloh and Muller, 1989). In the adult Hirudo nervous system, there is a single, unpaired S cell per midbody ganglion (Muller et ul., 1981). That cell sends two large axons into the median connective nerve, one anteriorly and the other posteriorly; these axons extend about halfway to the next anterior or posterior ganglion. The anterior axon makes an electrical synapse with the posterior axons of the next anterior S cell at this midway point, so that the ensemble of S cells forms a functionally syncytial chain along the ventral nerve cord (Muller and Carbonetto, 1979). T h e S cells in the anteriormost ganglia of Hirudo begin axonogenesis during E9-El0, as do other Hirudo neurons. By E12-El3, S cell axons from adjacent ganglia have met midway in the interganglionic connective nerves, and within a day of this meeting have become dye coupled. T h e embryonic S cell axons continue to elongate in the connective nerve for several days after this initial contact and coupling, until they overlap for as much as 76% of their length. Thus, the coupling of embryonic S cells does not inhibit their continued axonal extension (McGlade-McCulloch and Muller, 1989). By contrast, in adult specimens of H i n d u , S cell axons damaged by a crush of the connective nerve will regrow toward each other to reestablish their connection (Muller and Carbonetto, 1979), but axonal extension ceases as soon as dye coupling between anterior and posterior S cell axons has been reestablished (Scott and Muller, 1980).

4. Motor Neurons Most motor neurons project their axons across the ganglionic midline into the contralateral segmental nerve roots, on their way to the

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peripheral musculature. Unlike interneuron S or the mechanosensory neurons, adult motor neurons project no axons into anterior or posterior ganglia via the interganglionic connectives. Study of the morphogenesis of different types of motor neurons has revealed some significant differences in their developmental strategy. In Haementeria, motor neuron L (which forms extensive peripheral arborizations innervating the entire longitudinal musculature of the contralateral segmental body wall) begins axonogenesis early in stage 9, at about the same time as the other neurons of the ganglion (Kuwada, 1984). The L neuron projects a single axon, which grows across the midline into the contralateral posterior segmental nerve root, entering that root only after the axon of the mechanosensory P, neuron has done so. Growth cones of L axons are small, unlike the large and complex growth cones of P, and P, axons, which suggests that the growing L axons fasciculate with and follow the sensory axons in their peripheral outgrowth. Although each L neuron projects a short, anteriorly directed process into the contralateral neuropil of its ganglion, it produces no supernumerary axons that are later pruned or reabsorbed. Throughout development, the pattern of L axons is consistent with the adult pattern, as is the case for the axonal outgrowth pattern of vertebrate motor neurons (Lance-Jones and Landmesser, 1981; Tosney and Landmesser, 1985; Eisen et al., 1986). Further insights into the mechanism of L axon outgrowth were obtained from developmentally abnormal Haementeriu embryos in which the two halves of the germinal plate had failed to fuse, producing two separate haif-leeches with half-ganglia (Kuwada, 1984). In such specimens, the L axons could not follow their normal pathway across the ganglionic midline. Under these abnormal conditions, the L neuron produced several supernumerary axons that followed a diversity of abnormal pathways, such as into the ipsilateral segmental nerve roots and into the interganglionic connectives. This finding implies that, in response to abnormal developmental cues provided by the bisected germinal plate, the L motor neuron can generate more than a single axon and can send the supernumeraries along pathways not normally taken by its processes. L axons are, therefore, not absolutely constrained to follow the normal L pathways. In contrast to the single, definitive axon normally projected from the start by the L motor neuron of Haementeriu, the heart accessory (HA), annulus erector (AE), and “anterior pagoda” (AP) neurons of Himdo all generate supernumerary axons during their early development, axons that are lost in later development (Gao and Macagno, 1987a,b, 1988). At the outset of axonogenesis, all three types of neurons send axons across the ganglionic midline into both contralateral segmental nerve roots, as

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.. urn FIG. 18. HA neuron projection in Hirudo medzcinulis. Ganglionic outlines are indicated by dotted lines. (A) Normal projection pattern of mature HA neurons, consisting of an arborization within the neuropil and two major axons in the roots contralateral to the soma. Note that H A neurons normally lack axons in the interganglionic connective nerves. (B) Projection of H A neurons following the ablation of one embryonic H A neuron in MX. Central axons are maintained by the HA neurons that are ipsilateral to the ablated neuron in adjacent ganglia both anterior and posterior to the ganglion lacking one HA. Extra axons from M7 and M9 enter the periphery by way of the anterior and posterior nerve roots, in MX. The normal position of the ablated HA neuron is indicated by a circle. M7, M8, and M9 indicate ganglia in midbody segments 7, 8, and 9. (Data from Gao and Macagno, 1987b.)

well as into the anterior and posterior contralateral connective nerves. Later in development, the axons in the connectives regress, so that in adults the HA, AE, and AP neurons innervate only the periphery of their o w n segment and lack intersegmental projections (Fig. 18A). To determine whether interactions between ipsilateral intersegmental homologues play a role in this developmental loss of central axons, various neurons were ablated and the axonal projection patterns of neurons in adjacent segments were examined at later stages. Ablation of HA, AE, or AP neurons early in development enhanced the persistence of the central axons of their homologues in adjacent segments long after

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these axons normally would have disappeared (Fig. 18B) (Gao and Macagno, 1987a,b). This result is consistent with the hypothesis that transsegmental interactions among central axons of segmentally homologous neurons can lead to the elimination of interganglionic processes. However, severing the connection of AP and AE motor neurons to their peripheral muscle targets also enhanced the persistence of their central axons, suggesting that contact with their peripheral targets sensitizes these neurons to the central intersegmental interaction that promotes the elimination of their central processes (Gao and Macagno, 1988). 5. Other Effector Neurons Pruning of initially exuberant axonal outgrowth occurs also in the case of some Retzius neurons in Hirudo embryogenesis (Glover and Mason, 1986; Jellies et ul., 1987). These large, serotonergic neurons innervate muscles in the periphery and arborize extensively in the ganglionic neuropil. They are thought to have a modulatory effect on their targets (Lent, 1977; Mason and Kristan, 1982; Leake, 1986; Lent and Dickinson, 1989). In most segments, Retzius neurons project axons into the periphery by way of both ipsilateral segmental nerve roots and into the adjacent anterior and posterior ganglia by way of the ipsilateral connective nerves (Fig. 19A). The peripheral axons branch extensively within the muscles of the body wall. In the two reproductive segments, M5 and M6, however, the peripheral Retzius axons branch in the walls of the reproductive ducts peculiar to these segments, but do not extend into the muscle layers of the body wall. In the two reproductive segments, moreover, the Retzius neurons lack interganglionic axons (Fig. 19B). During the early phases of axonogenesis, all Retzius neurons in midbody ganglia extend axons into the periphery and toward adjacent ganglia via the interganglionic connectives. It is only at later developmental stages that the intersegmental processes extended by the Retzius neurons of segments M5 and M6 [labeled Rz(5) and Rz(6) in Fig. 201 stop growing and are eventually retracted. This loss of central processes is temporally correlated with the initiation of contact between the axons of Rz(5) and Rz(6) and their eventual peripheral targets, the reproductive ducts (Glover and Mason, 1986; Jellies et al., 1987). If interaction of Rz(5) and Rz(6) with their presumptive targets is prevented by ablating the embryonic reproductive ducts at about the time contact begins, the central axons of these neurons persist (Loer et ul., 1987). Thus, persistence of central processes of Rz(5) and Rz(6) neurons appears to depend upon the presence or absence of specific peripheral target tissue, as does the axonal pattern of motor neurons AE and AP. Unlike the case of the AE and AP neurons, however, intersegmental interactions among

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250pm FIG.19, Peripheral projection of Retzius neurons in standard and reproductive segments of H i r u h The outlines of ganglia are indicated by dotted lines. (A) Primary and secondary projection fields of Rz(9) in an embryo on E20. The projection reaches from the ventral midline (VM,dashed line) to the dorsal midline (DM, approximate position indicated by arrow) in three segments and consists largely of branches in an orthogonal pattern. (B) Projection of Kz(5) in an embryo on E20. Rz(5) lacks axons in the interganglionic connectives and branches densely around the male reproductive ducts, whose outline is indicated by dashed lines. (Unpublished data, K. A. French and W. B. Kristan, J r . )

central axons of segmental homologues appear to be unimportant in controlling the central morphology of Retzius neurons (Loer and Kristan, 1989~).

PERIPHERAL TARGETS F. INTERACTIONS BETWEEN NEURONSAND THEIR As in the developing vertebrate nervous system (Smith and Frank, 1987; Schotzinger and Landis, 1988), interactions between neurons and their peripheral targets play a role in leech neurogenesis (French and Kristan, 1992).These interactions have been examined most extensively

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in the reproductive segments M5 and M6 of Hirudo. The developmental interactions of Rz(5) and Rz(6) neurons with their target organs, the male and female reproductive ducts, affect not only the axonal projection pattern of these neurons, but also several other of their properties: soma size, complexity of neuropil arborization, synaptic inputs, and sign of cell membrane polarization in response to ACh (Fig. 20) (Jellies et al., 1987; Kristan and French, 1988; Loer and Kristan, 1989a; Wittenberg et al., 1990). For instance, during the early stages of axonogenesis, the morphology of Retzius neurons is indistinguishable in all midbody ganglia. But Rz(5) and Rz(6) begin to look different from their segmental homologues after their peripheral axons have approached the embryonic reproductive ducts (Glover and Mason, 1986; Jellies et al., 1987). If development is allowed to proceed normally, Rz(5) and Rz(6) develop smaller somata and sparser dendritic arborizations within the neuropil of their home ganglion, and they lack central intersegmental axons. Furthermore, stimulation of either a P mechanosensory neuron or of the neural circuit controlling swimming fails to evoke excitation of Rz(5) or Rz(6), whereas either stimulus is clearly excitatory for Retzius neurons in all other midbody ganglia. Finally, application of ACh to their somata hyperpolarizes Rz(5) and Rz(6), whereas all other Retzius neurons depolarize in response to applied ACh. Ablation of the embryonic reproductive ducts at the time when the peripheral Retzius axons would normally approach them causes Rz(5) and Rz(6) to develop a morphology (Loer et al., 1987; Loer and Kristan, 1989b), synaptic input patterns (Loer and Kristan, 1989a), and a response to ACh application (W. B. Kristan, Jr. and K. A. French, unpublished results) similar to that of standard Retzius neurons. The contact between embryonic reproductive ducts and growing axons of Rz(5) and Rz(6) generates immediate as well as long-term changes in their appearance. At the time Retzius axons first leave the ganglion, their growth cones are small and simple. Most Retzius growth cones remain simple, but within hours of reaching the reproductive ducts the growth cones of Rz(5) and Rz(6) expand, becoming both larger and more complex (French et al., 1992). When reproductive duct tissue is transplanted into nonreproductive segments, axons of the resident Retzius neurons approach the tissue and their growth cones enlarge, as they do in segments M5 and M6 (French et al., 1992). Although this early response to axonal contact with ectopic reproductive ducts in nonreproductive segments is similar to that normally seen in reproductive segments, it is not sufficient to confer the other morphological features of the Rz(5) and Rz(6) phenotype on Retzius neurons in segments other than M5 and M6 (Loer and Kristan, 1989~).

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The reproductive ducts affect the development of M5 and M6 in other ways, as well. In adults, M5 and M6 contain many more neurons than other midbody ganglia (Macagno, 1980), and most of these extra neurons are generated only near the end of embryogenesis and in postembryonic life (Stewart et al., 1986). However, in Hirudo there is at least one extra pair of neurons present solely in ganglion M6, the rostra1 penile evertor (RPE) cells (Zipser, 1979).These paired neurons are born and begin axonogenesis at about the same time as most other neurons, i.e., on E9 or E10. The presence of reproductive ducts has been implicated in both the generation of the extra cells in ganglia M5 and M6 and in shaping the final axonal pattern of the RPE neuron. If the reproductive ducts are ablated before E 16, no extra cells are subsequently added to the ganglia of M5 or M6. If the ducts are ablated on E 16 or thereafter, the extra cells appear on schedule (Baptista and Macagno, 1988a).Thus, the reproductive ducts influence segment-specific neurogenesis in these two ganglia (Baptista et al., 1990).This induction of extra neurons depends not only on a signal provided by the ducts, however, but also upon the competence of the neural precursor cells of ganglia M5 and M6 to respond to the signal; reproductive ducts transplanted ectopically into segments other than M5 and M6 fail to induce the production of extra neurons. Adult RPE neurons project axons to the contralateral periphery

FIG. 20. Features distinguishing Rz(5,6) from Rz(X) in Hirudo medicinalis. (A) Soma size and density of arborization within the neuropil (expressed as a percentage of the total neuropilar area containing processes of labeled neurons, as seen in camera lucida tracings). The soma size and the density of arborization increase much more rapidly between E9-11 and E20-24 in standard segments than they do in reproductive segments, causing Rz(5,6) to have distinctly smaller somata and smaller central arborizations by the end of embryogenesis. (Data from Jellies et al., 1987.) (B) Synaptic inputs onto Rz(X) lead to excitation when the tail is pinched (at times indicated by arrows in the left traces), when the swimcontrolling neural circuit is active (indicated by the horizontal bars in the middle traces, after tail pinches indicated by arrows), or when a pressure-sensitive mechanosensory neuron (P cell) in the same ganglion is stimulated by an intracellular electrode (at times indicated by the horizontal bars in the right traces). Rz(5,6) lack these inputs. The horizontal voltage calibration bar represents 2 mV for the third Rz(X) trace and for the second and third Rz(6) traces, 5 mV for the first Rz(6) trace, 10 mV for the first and second Rz(X) traces, and 20 mV for the P cell traces. The vertical time calibration bar indicates 1 sec for the first and second Rz(X) traces and for the first Rz(6) trace, 2 sec for the second Rz(6) trace, and 20 msec for the third traces for both Rz(X) and Rz(6). (Data from Loer and Kristan, 1989a.) (C) Response of Rz neurons to acetylcholine (ACh) ejected onto their soma in ganglia from which the glial sheath has been removed. Rz(X) neurons depolarize in response to ACh, whereas Rz(5,6) hyperpolarize. (Unpublished data, K. A. French and W. B. Kristan, Jr.)

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through the anterior segmental nerve root of ganglion M6, as well as via an interganglionic collateral process through the posterior segmental nerve root of ganglion M5 (Baptista and Macagno, 1988b). The reproductive ducts of segments M5 and M 6 are among the peripheral targets of the RPE axons. Early in axonogenesis, however, the RPE neurons send axons through both the anterior and posterior segmental nerve roots of ganglion M6 and both anteriorly and posteriorly into the interganglionic connective nerves. The supernumerary axons are pruned slowly in late embryonic life and possibly persist into postembryonic life. If contact between the RPE axons and the reproductive ducts is averted or disrupted, either by ablating the ducts or by severing the segmental nerves containing the RPE axons, the supernumerary axons persist (Baptista and Macagno, 1988b). T h e reproductive ducts thus seem to play an important role in shaping the definitive axonal projection pattern of RPE neurons. Thus, interactions between the embryonic reproductive ducts and neurons in segments M5 and M6 modify several disparate aspects of neuronal development, including neurogenesis, axonal growth and maintenance, synaptogenesis, and the production of neurotransmitter receptors. In some instances (e.g., in the differentiation of the Retzius, AE, and HA neurons) the interaction requires contact between peripheral processes and their targets, whereas in other cases (e.g., in the genesis of extra neurons) a diffusible factor might mediate the interaction.

G. NEURON-NEURON INTERACTIONS: THEORIGINS OF UNPAIRED NEURONS

Initial overproduction of neurons that subsequently compete with one another for survival is a prominent feature of vertebrate neurogenesis, wherein it serves to match neuronal populations to the available targets (Oppenheim, 1991; Truman, 1984). Although such overproduction is less prominent in the highly determinate development of the leech nervous system, the production of a dozen or so unpaired interneurons, in a typical segmental ganglion, from the initially symmetrical embryonic cell lineages of the leech nervous system offers several examples of competitive interactions that control neuronal survival or differentiation. Attention was first drawn to neuron-neuron competition in the leech when lineage tracers were used to determine the lineage of the unpaired, posteromedial serotonin neuron pms (Fig. 3). By double labeling

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embryos with lineage tracer and an antiserotonin antibody, it was shown that the pms neurons are derived from the N teloblast, like all other serotonin neurons. In some ganglia, pms arises from the right N teloblast and in other ganglia pms arises from the left (Blair and Stuart, 1982; Stuart et al., 1987), with the side of origin in a given segment varying randomly from specimen to specimen (Blair and Stuart, 1982; Stuart et al., 1987; Macagno and Stewart, 1987). Thus, each N teloblast gives rise to approximately half of the pms neurons in a normal embryo, but which half is indeterminate, so it was surprising when ablation of one N teloblast caused no deficit of pms neurons (Stuart et al., 1987). This result was explained by the observation that two pms neurons arise in each ganglion, one descended from each N teloblast. In a normal embryo, one o r the other pms dies during stage 10, but in an embryo deprived of one N teloblast, all of the pms neurons descended from the remaining N teloblast survive. Evidently, competition between pms cells derived from the two N teloblasts determines which pms neuron in a given ganglion survives and which dies (Stuart et al., 1987; Macagno and Stewart, 1987). T h e developmental mechanisms for producing two other unpaired neurons seem to be similar. Lineage-tracing studies indicate that the unpaired neurons pz4 (Figs. 9 and 10) and mz4 (Fig. lo), which descend from the P and M teloblasts, respectively, also arise from bilaterally paired precursors. One precursor in each segment dies near the end of stage 9 (Kramer and Weisblat, 1985; Shankland and Martindale, 1989; D. K. Stuart, S. M. Shankland, and S. A. Torrence, unpublished observations). Two other unpaired neurons represent a variation on this theme. These cells are the large rostra1 and caudal alternating SCP neurons (RAS and CAS; collectively called AS neurons) (Fig. 21A) (Blair et al., 1990). AS neurons, which stain intensely with an antibody against small cardiac peptide, are found on only the left or right side of any ganglion. Their development has been studied in glossiphoniid leeches. RAS neurons, which were shown to descend from the N teloblasts, are found only in ganglia Ml-M3 and in the fourth subesophageal ganglion. CAS neurons, which descend from the M teloblasts, are found only in ganglia M 18-M2 1 and in caudal ganglia. Each asymmetrically placed immunoreactive AS neuron seen in an adult nerve cord arises as one of a bilateral pair of cells that begin to stain with the anti-SCP antibody in late stage 10 o r early stage 1 1 . Subsequently, the cell that becomes the AS neuron maintains its immunoreactivity, while the other ceases to stain. T h e development of AS neurons differs from that of pms, pz4, or mz4 in that no evidence of cell death has been found, and it seems likely that

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Subesophageal Ganalion

“

B CAS

RAS 72%

71%

73%

78%

Abdominal Ganglia

41

15

65

18

CAS

Neuron Region

Tail Ganglion w

FIG. 21. Phenotypic specification of neurons by interactions within a segment and between segments. (A) Segmental distribution of SCP-immunoreactive KAS and GAS neurons (black dots) in the nerve cord of Helobdella triserialir. (B) Effect of unilateral RAS or CAS precursor ablations on the asymmetry of homologous AS neuron differentiation in adjacent segments. Ablation of a single AS neuron precursor (at a site indicated by X) caused the contralateral AS precursor (stippled) to express the mature AS phenotype of SCP-like immunoreactivity. The numbers in the left and right sides of the adjacent ganglia are the numbers of experimental embryos in which an unpaired, immunoreactive AS neuron was observed in those locations. There was a strong tendency for AS neurons in adjacent ganglia to alternate sides with the AS neuron in the lesioned ganglion. The percentage of alternation is indicated in italics beside the ganglia. (From Blair el al., 1990.)

the AS homologue survives but expresses an alternative neurochemical phenotype (Shankland and Martindale, 1989).Like the survival of pms, the neurochemical phenotype of AS neurons is controlled by competition between contralateral homologues, because ablation of AS precursors on one side of appropriate segments causes the precursors on the other side invariably to take the AS (Le., SCP+) phenotype. By varying

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the timing of these ablations it was shown that the competitive interaction occurs after the terminal mitosis of the AS precursors (Martindale and Shankland, 1990). A striking feature of the distributions of many unpaired leech neurons is that the side of origin of a given neuron tends to alternate from one ganglion to the next along the nerve cord (Fig. 21A). This has been shown in Hirudo for the pms neurons (Macagno and Stewart, 1987) and six types of unpaired SCP neurons, including the homologue to CAS (Evans and Calabrese, 1989), and in Theromyzon and Helobdella for mz4 and the As neurons (Shankland and Martindale, 1989). Such segmentto-segment alternation suggests that the outcome of competition between bilaterally homologous cells within a given segment is biased by interactions with cells in adjacent segments. This proposal was directly tested for the AS neurons by ablating, in a single hemiganglion, a small cluster of neurons that included the precursor of an AS neuron (Fig. 21B) (Blair et al., 1990). This manipulation caused the contralateral homologue of the ablated precursor to take the AS phenotype, confirming the intraganglionic interaction proposed above and imposing a particular sidedness to the AS symmetry in the operated segment. T h e AS neurons in most segments adjacent to an operated segment were found on the same side as the lesion-they alternated sides with the AS neuron in the operated segment. Thus, consistent with the proposed intersegmental interaction, experimentally imposed AS sidedness in one ganglion biased the outcomes of AS competitions in the adjacent ganglia. Many of the unpaired neurons whose side of origin tends to alternate from one segment to the next project axons to one or both adjacent ganglia via the interganglionic connective nerves. These axonal projections are proposed to mediate the interganglionic interaction (Macagno and Stewart, 1987; Shankland and Martindale, 1989; Martindale and Shankland, 1990; Blair et al., 1990). To test this proposal, the connective nerves between CAS-containing ganglia were transected in late stage 10 o r early stage 11 Theromyzon embryos. As predicted, such transection reduced the frequency of CAS alternation across the cut (Martindale and Shankland, 1990).

VI. Conclusions

As shown by the research summarized in this review, embryogenesis of the leech nervous system is highly determinate in the sense that during normal development the genealogical origin of each identified neu-

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ron can be traced, via a sequence of stereotyped cleavages, to the zygote. This determinacy might suggest that the developmental fate of a given cell is somehow governed entirely by its particular line of descent. However, cellular interactions strongly affect the fate taken on by at least some neural precursor cells, as, for example, is the case for the O/P kinship group. Thus, although the characteristic phenotype of each identified neuron stems from a series of increasingly restrictive developmental commitments made by its ancestors, and although for many neurons these developmental choices would seem to rely strongly on cell lineage, interactions with other cells cannot be ignored as a source of significant developmental information. I t seems to be generally thought that cell fates become assigned when the developmental possibilities of pluripotent cell lines are limited by a set of sequential commitment steps that end in differentiation into the final set of phenotypes emanating from each pluripotent cell line. It has been thought that such a process depends on a stepwise, typologically hierarchic sequence of commitments (Slack, 1983). For instance, a cholinergic (excitatory) motor neuron would arise in a leech along the following pathway: ( 1 ) from a cell clone committed to expressing characters that distinguish ectoderm from mesoderm, to a neuroectoderm subclone committed to expressing characters that distinguish nervous tissue from epidermis; (2) from the neuroectoderm subclone to a neuronal subclone committed to expressing characters that distinguish neurons from glia; (3) from the neuronal subclone to a motor neuron subclone committed to expressing characters that distinguish motor neurons from sensory neurons; and finally, (4)from the motor neuron subclone to a cholinergic subclone committed to expressing the gene that encodes cholineacetyltransferase (typical of excitatory motor neurons), rather than expressing the gene for glutamic acid decarboxylase (typical of inhibitory motor neurons). Contrary to this hypothetical pattern, during the development of the leech nervous system the sequence of commitment steps appears to be typologically arbitrary, rather than hierarchic. In fact, the phenotype of particular neurons shows little correlation with membership in any particular kinship group (cf. Fig. 10). Indeed, even after a primary blast cell has undergone one o r two divisions in the generation of its segmental founder clone, one of its daughter blast cells may still give rise to a mixed clone of neural and epidermal cells (Shankland and Stent, 1986). Hence in the leech, instead of the proposed pattern, the neurodevelopmental pathways seem to be mainly topographically hierarchic, rather than typologzcally hierarchic. In other words, it is the position of two cells, rather than their phenotype, that is usually correlated with their geneological closeness.

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Admittedly, the serotonin neurons, all being derived from the N teloblast, are genealogically more closely related to one another than they are to the dopamine neurons, which are derived from the OPQ blastomere. This relationship might suggest that the NOPQ blastomere cleaves in a typologically hierarchic fashion, according to which the N teloblast daughter is committed to generate serotonin neurons and the OPQ blastomere daughter is committed to generate dopamine neurons (cf. Figs. 3, 9, and 10). However, these two types of monoamine neurons account for only a small fraction of all the diverse progeny of the N and OPQ cells, so it appears more reasonable to invoke a topographic interpretation of the cleavage pattern: nearly all of the N kinship group cells (including the serotonin neurons) are destined for the CNS, whereas the bulk of the OPQ-derived cells (including the dopamine neurons) are destined for the body wall (Weisblat et aZ., 1984). The topographic rationale can be extended further to the cleavage pattern of the OPQ blastomere, whose Q and OP daughter cells primarily generate descendants in the dorsal and ventral body wall, respectively. Nevertheless, the topographic hierarchy in the cleavage pattern is far from absolute. Some descendants of the Q and of the OP blastomeres are destined for the central nervous system on the ventral midline, whither they migrate from the lateral germinal plate, where the cleavage pattern initially places them. Thus, topography based on lineage pattern may exert a strong influence upon the choice of developmental fates among leech neurons, but many neurons are clearly not restricted to their original location. Two kinds of agents are commonly thought to contribute to developmental commitment under either the typologically or the topographically hierarchic mode. One of these agents consists of a set of intracellular determinants, which would account for the differential commitment of sister cells in terms of unequal partition of cellular elements in successive cell divisions. For instance, a pluripotent cell might possess two determinants, a and 6, that are necessary for producing cell types A and B respectively. Commitment of a daughter cell to fate A (and loss of pluripotency) would occur at an asymmetric cell division in which the daughter cell received only a and not b. Under this mechanism, cell lineage would play a governing role in cell commitment by consigning particular subsets of intracellular determinants to particular cells. The other commonly considered agent leading to commitment consists of a set of intercellular inducers whose elements are anisotropically distributed through the volume of the embryo. In this construction, a pluripotent cell would be one that was capable of responding to either of two inducers, a or 6, that were necessary for producing cell types A and B, respectively. Commitment of a cell to fate A (and the loss of pluripotency)

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would occur when the cell responded to inducer a at some crucial stage of development and lost the ability to respond to 6. Under this mechanism, cell lineage could play a crucial role in cell commitment by placing particular cells at particular sites within the inductive field, hence governing the pattern of their exposure to inducers. (Note that this idea leaves the source of the competence to respond to a and b open to conjecture.) Developmental studies carried out with nematodes (Laufer et al., 1982) and ascidians (Whittaker et al., 1977) have shown that cell lineage can play a governing role in cell commitment by bringing about the orderly, unequal partitioning of intracellular determinants among daughter cells in successive cell divisions. T h e mitotic partition of teloplasm during the very earliest leech embryonic cleavages appears, similarly, to exert a strong influence on the subsequent development of leech blastomeres A, B, C, and D (Astrow et al., 1987; B. Nelson, personal communication). It is also clear, however, that in the determinate development of the leech, the orderly topographic placement of cells relative to anisotropically distributed intercellular inducers must play a major role. This conclusion follows from the numerous cases of developmentally critical cellular interactions that are summarized in this review. Such interactions are important during early development, as in the potential transfating of primary O/P blast cells, and later when neuronal phenotype depends on contacting other neurons or nonneuronal tissue. Either of these two classes of agents, or perhaps a combination of the two, may determine the identity of particular leech neurons. However, the stereotyped patterns of migration and of specific process outgrowth that characterize many identified neurons suggest that the ability to recognize and to respond appropriately to particular signals in the environment plays an important role in shaping the details of connectivity within the leech nervous system. Migration and process outgrowth must depend upon intercellular signals, because manipulations that disrupt normal interactions but leave precursors of the identified neurons intact disrupt the precision of these patterns. The competence to respond to such external cues might originate either from particular intracellular agents or from a previous commitment response to intercellular agents. For example, the commitment to follow a particular migratory path apparently arises before central neurons derived from the OPQ clones begin their journey, and it could, thus, be the product of either type of agent. In contrast, fine tuning the peripheral and central processes of many neurons occurs long after the cells become postmitotic and depends upon the specific interactions to determine a final phenotype. 'These developmental events must rely upon intercellular agents, be-

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cause in the absence of the specific targets the final phenotype of the neuron is abnormal. Intracellular agents are insufficient, on their own, to generate the entire ensemble of normal characteristics in these cells. As described in this review, studies carried out over the past decade have produced a reasonably good understanding of the genesis of the leech nervous system, in many of its structural and some of its functional aspects, and at both the cellular and intercellular levels. There has recently been an explosive development of novel techniques for analyzing systems at the molecular biological level, and it seems likely that these techniques offer an excellent opportunity to understand the development of leeches-and hence the development of other organisms, as well-at a more detailed level than has heretofore been possible. Identified cells in leech embryos are readily accessible to various kinds of biochemical manipulations, exemplified by the injection of macromolecules, and leech embryos offer the unusual opportunity to follow animal development at the cellular level from uncleaved egg to mature tissue in embryos large enough to allow molecular and surgical manipulations. Thus, it is likely that identifying the molecular nature of both intracellular determinants and intercellular inducers will be one of the main items on the agenda for future research concerning the development of the leech nervous system. A second item on that agenda is likely to focus on the behavioral effects of the leech embryo’s increasingly complex nervous system. A major function of nervous systems is to generate behavior appropriate to conditions within the environment, and the complexity and precision with which elements in the system connect to one another are thought to determine the complexity and precision of the behavioral responses that are possible. Embryonic leeches begin behaving early, and this behavior has been found to differ, in several ways that are described in this review, from that in the adult. Correlating the increasing complexity and interconnected nature of the nervous system in embryonic leeches with their increasingly complex and precise behavior should teach us more about how the nervous system acts as the biological substrate for behavior.

References

Anderson, D. T. (1973). “Embryology and Phylogeny in Annelids and Arthropods.” Pergarnon, Oxford. Apathy, S. (1889). Biol. Centralbl. 9, 600-608. Astrow, S., Holton, B., and Weisblat, D. (1987).Dev.Biol. 120, 270-283.

188

CXNTHER S. STENT et al.

Baader, A,, and Kristan, W. B., Jr. (1990). In “Brain, Perception and Recognition” (N. Elsner and G. Roth, eds.), p. 60. Thierne, New York. Baptista, C. A , , and Macagno, E. R. (1988a).J. Neurobiol. 19, 707-726. Baptista, C. A., and Macagno, E. R, (1988b). Neuron 1, 949-962. Baptista, <;. A., Cxrshon, T. R., and Macagno, E. R. (1990). Nature (London) 346,855-858. Baylor, D. A.,and Nicholls, J. G . (1969).J. Physiol. (London) 203, 591-609. Belanger. J. H., and Orchard, I. (1986). Brain Res. 382, 387-391. Belanger. J. H.. and Orchard, I. (1988).J. Comp. Physiol. A 162A, 408-412. Belarderti, F., Biodi, C., Colornbaioni, L., Brunelli, M., and Trevisani, A. (1982). Brain Res. 246, 89-103. Bergh, R. S. (1885). Arb. Znol. Imt. U’urzburg 7, 231-291. Bissen, S. T., and Weisblat, D. A. (1987).J. Neurobiol. 18, 251-269. Bissen, S. T., and Weisblat, D. A. (1989). Development (Cumbridge. UK) 106, 105-1 18. Blackshaw, S. E. (1981a).J. Physiol. (London) 317, 81P-82P. Blackshaw, S. E. (1981b).J. Physiol. (Lordoti) 320, 219-228. Blackshaw, S. E. (1981~). In ”Neurobiology of the Leech,” (K. J. Muller, J. G. Nicholls, and G. S. Stent, eds.), pp. 113-146. Cold Spring Harbor Lab., Cold Spring Harbor, New York. Blackshaw. S., and Nicholls, J. (1979).J. Physzol. (London) 292, 26-27. Blair, S. S. (1982). Dai. Riot. 89, 389-396. Blair. S. S. (1983). Drv. Rial. 95, 65-72. Blair, S. S., and Stuart, D. K. (1982). Soc. Neurosci. Abstr. 8, 16. Blair. S. S., and WeisbIat, D. A. (1982). Dev. Biol. 91, 64-72. Blair, S. S., and Weisblat, D. A. (1984). Dev. B i d . 101, 318-325. Blair, S. S.. Martindale, M. Q.,and Shankland, M. (1990).J. Neurosci. 10, 3183-3193. Boulis, N. M., and Sahley, <;. L. (1988).J. Neurosci. 8, 4621-4627. Braun,.J., and Stent, G. S. (1989a). Dm. B i d . 132, 471-485. Braun, J . , and Stent, G. S. (1989b). Drv. B i d . 132, 486-501. Brodfuehrer, P. D., and Cohen, A. €1. (199O).J. Exp. Biol. 154, 567-572. Brodfuehrer, P. D., and Friesen, W. 0. (1986).J. Comp. Pliysiol. A 159A, 489-502. Bullock, T. H., and Horridge, G. A. (1965). “Structure and Function of the Nervous System of Invertebrates.“ Freeman, Sail Francisco, California. Buznikov, G. A., Chudakova, L. V., Berdychwa, L. V., and Vyazrnina, N. M. (1968).Embtgol. Exp. Morphol. 20, 1 19- 128. Calabrese, R. L., and Arbas, E. A. (1989). I n “Neuronal and Cellular Oscillators” (J. W. Jacklet, ed.), pp. 237-267. Dekker, New York. Calabrese, R. L., and Peterson, E. (1983). Symp. Sor. Exp. Biol. 37, 195-221. Calabrese, R. L., Angstadt, J. D., and Arbas, E. A. (1989). I n “Perspectives in Neural Systems and Behavior” (T. J. Carew and D. B. Kelley, eds.), pp. 33-50, Liss, New York. Caudy, M., and Bentley, D. (1986).J. Neurosci. 6, 364-379. Cline, H. T. (1983).J. Conrp. Neurol. 215, 351-358. Cline, H. T. (1986).J . iVeurusci. 6,2848-28.56. Coggeshall, R. E., and Fawcett, D. W. (1964).J. Neurpltysiosiol. 27, 229-289. Crick, F. H. C;., and Lawrence, P. A. (197.5). Science 189, 340-347. Dawydoff, C:. (1959). [ti “Traite de Zoologie” (P.-P. Grasse, ed.), Vol. 5, pp. 594-686. Masson, Paris. Dehski, E. A., and Friesen, W. 0. (1986).J. Neurophysiol. 55, 977-994. Deppe, F.Schierenberg, E., Cole, T., Krieg, C., Schrnitt. D., Yoder, B., and von Ehrenstein, G. (1978). Proc. Nutl. Acad. Sci. U.S.A. 75, 376-380. Derosa, Y . S., and Friesen, W. 0. (1981). B i d . Bull. (Woo& Hole, Mass.) 160, 383-393. Dietzel, 1. D., and Gottmann, K. (1988). Dev. Biol. 128, 277-284.

LEECH NERVOUS SYSTEM DEVELOPMENT

189

Eisen, J. S., Myers, P. A., and Westerfield, M. (1986). Nature (London) 320, 269-271. Evans, B. D., and Calabrese, R. L. (1989). Cell Tissue Res. 257, 187-199. Evans, B. D., Pohl, J., and Calabrese, R. L. (1990). SOC.Neurosci. Abstr. 16, 306. Fernandez, J. (1980). Dev. Biol. 76, 245-262. Fernandez, J., and Olea, N. (1982). In “Developmental Biology of Freshwater Invertebrates” (F. W. Harrison and R. R. Cowen, eds.), pp. 317-361. Liss, New York. Fernandez, J., and Stent, G. S. (1980). Dev. Biol. 78, 407-434. Fernandez, J., and Stent, G. S. (1983).J . Embryol. Exp. Morphol. 72, 71-96. Fitzpatrick-McElligott, S., and Stent, G. S. (1981).J. Neurosci. 1, 901-907. Frank, E., Jansen, J. K. S., and Rinvik, E. (1975).J. Comp. Neurol. 159, 1-13. French, K. A., and Kristan, W. B., J . (1992). I n “Determinants of Neuronal Identity” (M. Shankland and E. Macagno, eds.), Academic Press, San Diego, California (in press). French, K. A., Jordan, S., Loer, C., and Kristan, W. B., Jr. (1992). Dev. Biol. (in press). Friesen, W. 0. (1981).J. Exp. Biol. 92, 255-275. Friesen, W. 0. (1985).J. Comp. Physiol. A 156A, 231-242. Friesen, W. 0. (1989). I n “Neuronal and Cellular Oscillators”(J. W. Jacklet, ed.), pp. 269316. Dekker, New York. Friesen, W. O., and Stent, G. S. (1977). Biol. Cybernet. 28, 27-40. Friesen, W. O., Poon, M., and Stent, G. S. (1978).J. Exp. Biol. 75, 25-43. Gao, W.-Q., and Macagno, E. R. (1987a).J. Neurobiol. 18, 43-59. Gao, W.-Q., and Macagno, E. R. (1987b).J. Neurobiol. 18, 295-313. Gao, W.-Q., and Macagno, E. R. (1988). Neuron 1,269-277. Garcia-Bellido, A., and Merriam, J. R. (1969).J. Exp. Zool. 170, 61-76. Garcia-Bellido, A., Ripoll, P., and Morata, G. (1973). Nature (London), New Biol. 245,251253. Gee, W. (1913). Univ. C a l f . Berkelq, Publ. Zool. 11, 197-305. Gimlich, R. L., and Braun, J. (1985). Dev. Biol. 109, 509-514. Glover, J. C., and Kramer, A. P. (1982). Science 216, 317-319. Glover, J. C., and Mason, A. (1986). Dev. Biol. 115,256-260. Glover, J. C., and Stuart, D. K. (1983). SOC.Neurosci. Abstr. 9, 606. Glover, J. C., Stuart, D. K., Cline, H. T., McCaman, R. E., Magill, C., and Stent, G. S. (1987).J. Neurosci. 7, 581-594. Goodman, C. S., and Spitzer, N. C. (1979). Nature (London) 280, 208-214. Gray, J. (1968). “Animal Locomotion.” Weidenfels & Nicholson, London. Gray,J., Lissman, H. W., and Pumphrey, R. J. (1938).J. Ex$. Biol. 15, 408-430. Harant, H., and Grass6, P.-P. (1959). In “Trait6 de Zoologie” (P.-P. Grasse, ed.), Vol. 5, pp. 471-593. Masson, Paris. Haydon, P. G., McCobb, D. P., and Kater, S. B. (1984). Science 226, 561-564. Henderson, T. B., and Strong, P. N., Jr. (1972). Cond. Reflex 7, 210-215. Ho, R. K., and Weisblat, D. A. (1987). Dev. Biol. 120, 520-534. Ho, R. K., Ball, E. E., and Goodman, C. S. (1983).Nature (London) 301, 66-69. Jellies, J. (1990). Trends Neurosci. 13, 126-131. Jellies, J., and Kristan, W. B., Jr. (1988a).J. Neurosci. 8, 3317-3326. Jellies, J., and Kristan, W. B., Jr. (1988b). J. Neurobiol. 19, 153-165. Jellies, J., Loer, C. M., and Kristan, W. B., Jr. (1987).J. Neurosci. 7, 2618-2629. Johansen, J., Hockfield, S., and McKay, R. D. G. (1984). J. Comp. Neurol. 226, 263-273. Karrer, T., and Sahley, C. L. (1988). Behav. Neural Biol. 50, 311-324. Keleher, G. P., and Stent, G. S. (1990). Proc. Natl. Acad. Sci. U.S.A. 87, 8457-8461. Kramer, A. P., and Goldman, J. R. (198l).J. Comp. Physiol. 144, 435-448. Kramer, A. P., and Kuwada, J. Y.(1983).J. Neurosci. 3, 2474-2486. Kramer, A. P., and Stent, G. S. (1985).J. Neurosci. 5, 768-776.

190

GC‘NTHER S. S T E N T et al.

Kramer, A. P., and Stuart, D. K. (1982). Soc. Nwrosci. Abstr. 8, 16. Kramer, A. P., and Weisblat, D. A. (1985).J . Neurosci. 5, 388-407. Kramer, A. P., Goldman, J. R.,and Stent, G. S. (1985).J. Neurosci. 5 , 759-768. Kretz, J. R.,Stent, G. S., and Kristan, W. B., Jr. (1976).J. Comp.Physiol. 106, 1-37. Kristan, W. B., Jr. (1982).j. Exp. Bzol. 96, 161-180. Kristan, W.B., Jr., and Calabrese, R. L. (1976).J. Exp. Bzol. 65, 643-668. Kristan, W.B., Jr., and French, K. A. (1988). SOC.Neurosci. Abstr. 14, 164. Kristan, W.B., Jr., and Nusbaum, M. P. (1983). J. Physiol. (Paris) 78, 743-747. Kristan, W.B., Jr.. and Weeks, J. C. (1983). Sywip. SOC.Exp. Biol. 39, 243-260. Kristan, W. B., Jr., Stent, G. S., and Ort, C. A. (1974a).J. Comp. Physiol. 94, 97-1 19. Kristan, W. B., Jr., Stent, G. S., and Ort, C. A. (1974b).J. Comp.Pliysiul. 94, 155-176. Kristan, W. B., Jr., McGirr, S. J., and Simpson, G. V. (1982).J. Exp. B i d . 96, 143-160. Kristan, W. B., Jr., Weisblat, D. A., and Radocic, T. (1984). In “Invertebrate Models in Aging Research” (D. H. Mitchell and T. E. Johnson, eds.), pp. 95-119. CRC Press, Boca Raton, Florida. Kristan, W.B., Jr., Wittenberg, G., Nusbaum, M. P., and Stern-Tomlinson, W. (1988). In “Invertebrate Neuroethology” (J. Camhi, ed.), pp. 383-389. Birkhaueser Verlag, Basel. Kuffler, D. P. (1978).J. Comp.Physiol. A l24A, 333-338. Kuffler, D. P., and Muller, K. J. (1974).J. Neurobiol. 5, 331-348. KufRer, S. W.,and Potter, D. D. (1964).J. Neurophysiol. 27, 290-320. Kuhlman, J. R., Li, C., and Calabrese, R. L. (1985).J. Neurosci. 5, 2301-2309. Kusano, K.,Miledi, R., and Stinnakre, J. (1977). Nature (London) 207, 739-741. Kuwada, J. Y. (1982). 1n “Neuronal Development: Cellular Approaches in Invertebrates” (C. Goodman and K. Pearson, eds.), pp. 877-881. MIT Press, Cambridge, Massachusetts. Kuwada, J. Y. (1984).J . Embtyol. Exp. Murphol. 79, 125-137. Kuwada, J. Y., and Kramer, A. P. (1983).J. Neurosci. 3, 2098-21 11. Lance-Jones, C., and Landmesser, L. T. (1981). Proc. R . SOC.Londuu, S m B 214, 1-18. Lauder. J. M., and Krebs, H. (1978). Dev. Neurosci. 1, 15-30. Laufer, J., Bazzicalupo, P., and Wood, W. B. (1982). Nature (London) 297, 584-587. Leake, L. D. (1986).Comp. Biochem. Physiol. C 83C, 229-239. Le Douarin, N. (1973). Dm.Biol. 30, 217-222. Le Douarin, N. (1984). “The Neural Crest.” Cambridge Univ. Press, New York. Lent, C. M. (1973). Science 179,693-696. Lent, C . M. (1977). Prug. Ivmrubiol. 8, 81-117. Lent, C. M. (1982). Histochemisl~75, 77-89. Lent, C. M., and Dickinson, M. H. (1989). Am. Zool. 29, 1241-1254. Lent, C.M., Ono, J., Keyser, K. T., and Karten, H. J. (1979).J. Neurochem. 32, 1559-1563. Lent, C. M.,Fliegner, K. H., Freedman, E., and Dickinson, M. H. (1988).J. E x f . Biol. 137, 5 13-527. Levine, M., and Macagno, E. (1990).,4n?iu. Rpts Neurosci. 13, 195-225. Li, C.,and Calabrese, R. L. (1985). J. Comp. Neurol. 232, 414-424. Lockery, S. R., and Kristan, W. B., Jr. (1990a).J. Neurosci. 10, 1811-1815. Lockery, S. R., and Kristan, W. B., Jr. (1990b).J. Neurosci. 10, 1816-1829. Lockery, S. R.,and Kristan, W. B., Jr. (1991).J. Comp. Physiol. A 168A, 165-177. Lockery, S. R.,Wittenberg, G., Kristan, W. B., Jr., and Cottrell, G. W. (1989). Nature (London) 540, 468-47 1. Loer, C. M., and Kristan, W. B., Jr. (1989,). Science 244, 64-70. Loer, C.M., and Kristan, W. B., J . (1989b).J. Neurosci. 9, 513-527. Loer, C. M., and Kristan, W. B., J . (1989~).J. Neurosci. 9, 528-538.

LEECH NERVOUS SYSTEM DEVELOPMENT

191

Loer, C. M., Jellies, J., and Kristan, W. B., Jr. (1987).J. Neurosci. 7, 2630-2638. Macagno, E. R. (1980).J. Comp. Neurol. 190,283-302. Macagno, E. R., and Stewart, R. R. (1987).J. Neurosci. 7, 1911-1918. Magni, F., and Pellegrino, M. (1978).J. Comp. Physiol. 124,339-351. Mann, K. H. (1962). “Leeches (Hirudinea).” Pergamon, Oxford. Martindale, M. Q., and Shankland, M. (1988). Dev. Biol. 125,290-300. Martindale, M. Q., and Shankland, M. (1990). Dm. Biol. 139,210-226. Mason, A., and Kristan, W. B., Jr. (1982).J. Comp. Physiol. A 146A, 527-536. Mason, A., Sunderland, A. J., and Leake, L. D. (1979). Comp. Biochem. Physiol. C 63C,359361. McGlade-McCulloh, E., and Muller, K. J. (1989). Neuron 2, 1063-1068. Miller, J. P., and Selverston, A. I. (1979). Science 206, 702-704. Mintz, B. (1965). Science 148, 1232-1233. Muller, K. J. (1979). Biol. Rev. Cambridge Philos. Soc. 54, 99-134. Muller, K. J. (1981). In “Neurobiology of the Leech (K. J. Muller, J. G. Nicholls, and G. S. Stent, eds.), pp. 79-1 11. Cold Spring Harbor Lab., Cold Spring Harbor, New York. Muller, K. J., and Carbonetto, S. (1979).J. Comp. Neurol. 185,485-516. Muller, K.J., and McMahan, U. J. (1976). Proc. R. SOC.London, Ser. B 194,481-499. Muller, K. J., and Scott, S. A. (1981).J. Physiol. (London) 31, 740-756. Muller, K. J., Nicholls, J. G., and Stent, G. S., eds. (1981). “Neurobiology of the Leech.” Cold Spring Harbor Lab., Cold Spring Harbor, New York. Nicholls, J. G., and Baylor, D. A. (1968).J. Neurophysiol. 31, 740-756. Nicholls, J. G., and Purves, D. (1970).J. Physiol. (London) 209, 647-667. Nicholls, J. G., and Purves, D. (1972).J. Physiol. (London) 225, 637-656. Nicholls, J. G., and Van Essen, D. (1974). Sci. Am. 230, 38-48. Norris, B. J., and Calabrese, R. L. (199O).J.Comp. Physiol. 16, 211-214. Nusbaum, M. P., and Kristan, W. B., Jr. (1986).J. Exp. Biol. 122, 277-302. Oppenheim, R. W. (1991). Annu. Rev.Neurosci. 14,453-502. Ort, C.A., Kristan, W. B., Jr., and Stent, G. S. (1974).J. Comp. Physiol. 94, 121-154. Phillips, C. E., and Friesen, W. 0. (1982).J. Neurobiol. 13, 473-486. Poon, M., Friesen, W. O., and Stent, G. S. (1978).J. Exp. B i d . 75, 45-63. Rakic, P. (1972).J. Comp. Neurol. 145,61-84. Ram6n y Cajal, S. (1904). Trab. Lab. Invest. B i d . Univ. Madrid 3, 287-297. Ram6n y Cajiil, S. (1929). “Studies on Vertebrate Neurogenesis” (L. Guth, trans.). (Reprint, Thomas, Springfield, Illinois, 1960). Retzius, G. (1891). Biol. Unters. [N.S.] 2, 1-28. Reynolds, S. A., and Kristan, W. B., Jr. (1989). Soc. Neurosci. Abstr. 15, 349. Rude, S. (1969).J. Comp. Neurol. 136, 349-371. Sahley, C. L., and Ready, D. F. (1988).J. Neurosci. 8, 4612-4620. Sandig, M., and Dohle, W. (1988).J. Morphol. 196,217-252. Sargent, P. B. (1977).J. Neurophysiol. 40, 453-460. Sawyer, R. T. (1972). “North American Fresh Water Leeches, Exclusive of the Piscicolidae, with a Key to All Species.” Univ. of Illinois Press, Urbana. Sawyer, R. T. (1981). In “Neurobiology of the Leech” (K. J. Muller, J. G. Nicholls, and G. S. Stent, eds.), pp. 7-26. Cold Spring Harbor Lab., Cold Spring Harbor, New York. Sawyer, R. T. (1986). “Leech Biology and Behaviour,” Vols. 1,2, and 3. Oxford Univ. Press (Clarendon), London and New York. Sawyer, R. T., LePont, F., Stuart, D. K., and Kramer, A. P. (1981). B i d . Bull. (WoodF Hole, Mus.) 160,322-33 1. Schleip, W. (1936). I n “Klassen und Ordnungen des Tierreichs” (H. G. Bronn, ed.), Vol. 4, Div. 111, Book 4, Part 2, pp. 1-121. Akad. Verlagsges., Leipzig.

192

CXNTHER S. STENT et 01.

Schotzinger, R. J., and Landis, S. C. (1988). Nature (London) 335, 637-639. Schousbve, A . (1981). Int. Rev. iveurobiol. 22, 1-45. Scott, S. A.. and Muller, K. J . (1980). Dezj. Bzol. 80, 345-363. Shankland, M. (1984). iVature (London) 307, 541-543. Shankland, M. (1987a). Deil. B i d . 123, 85-96. Shankland, XI. (1987b). Drv. B i d . 123, 97-107. Shankland, M., and Martindale, hl. Q. (1989). Deu. B i d . 135, 431-448. Shankland, M., and Stent. G. S. (1986). In “Genes, Molecules and Evolution” 0. P. Gustafson, <;. L. Stebbins, and F. J. Ayah, eds.), pp. 21 1-233. Academic Press, Orlando, Florida. Shankland, M., and Weisblat, D. A. (1984). Deu. Bzol. 106, 326-342. Slack, J. M. W. (1983). “From Egg to Embryo.” Cambridge Univ. Press, London and New York. Smith, C. L., and Frank, E. (1987).J. X’eurosci. 7, 1537-1549. Stent, G . S. { 1985). Philos. Trarw. R. Sac. London, Ser. B 312, 3- 19. Stent, G. S., and Kristan, W. B. (1981). I n “Neurobiology of the Leech” (K. J. Muller, J. G. Nicholls, and C. S. Stent, eds.). pp. 113-146. Cold Spring Harbor Lab., Cold Spring Harbor, N e w York. Stem, <.; S., Kristan, W. B., Jr., Friesen, MI. O., Ort, C. A,, Poon, M., and Calabrese, R. L. (1978). Science 200, 1348-1357. Stent, G . S., Thompson, \\’.J., and Calabrese, R. L. (1979). Physiol. Rev. 59, 101-136. Stent, G. S., Weisblat, D. ii., Blair, S. S., and Zackson, S. L. (1982). In “Neuronal Development” (N. Spitzer, ed.), pp. 1-44. Plenum, New York. Stern, C . (1968). “Genetic Mosaics and Other Essays.” Harvard Univ. Press, Boston, Massachusetts. Stern-Tonilinson. W., Nusbaum, M. P., Perez, L. E., and Kristan, W. B., Jr. (1986).J.Comp. Phyiol. 158, 593-603. Stewart, R. R.,Spergel, D., and Macagno, E. R. (1986).J. Comp. Neurol. 253, 253-259. Stewart, R. R.,Gao, W.-Q., Peinado, A,, Zipser, B., and Macagno, E. R. ( 1 987).J. Neurosci. 7, 1919-1927. Stewart, W7.b’. (1978). Cell (Camhidge, Mass.) 14, 741-759. Stewart, W. W.( 198 1). .Vulure (London) 292, 17-2 1. Stuart, A. E. (1970).J. Pltysiol. (London) 209, 627-646. Stuart, A. E., Hudspeth, A. J., and Hall Z. W. (1974). Cell Tissue Re$. 153, 55-61. Stuart, 1). K., Thompson, J., Weisblat, D. A., and Kramer, A. (1982).Soc. Neurosci. Abstr. 8, 1.5. Stuart, D. K., Blair, S. S., and Weisblat, D. A. (1987).J. Neurosci. 7, 1107-1 122. Stuart, D. K., Torrence, S. A,, and Law, M. I. (1989).Deu. Biol. 136, 17-39. Sturtevant. A . H. (1929). 2. Wks. 2001.135, 325-356. Sulston, J. E., and Horvitz, H. R. (1977). Dev. Biol. 56, 110-156. Sulston, J. E.. and Horvitz, €3. R. (1981). Deu. Biol. 83, 41-55. Sulston, J. E., and White, J. G. (1980). Deu. B i d . 78, 577-597. Sulston, J. E., Schierenkrg, E., White, J. G., and Thomson, J. N. (1983).Dev. Bzol. 100,64119.

Tarkowski, A. K. (1961). Nature (London) 190, 857-860. IFnnyson, V. M . (1965).J . Couip. Neurol. 124, 267-318. Thompson. M! J., and Stent, G. S. (1976a).J. Comp. Physiol. 111, 261-279. Thompson, W. J., and Stent, G. S. (1976b).J . Comp. Physiol. 111, 281-307. Thompson, W. J., and Stent, G. S. (1976c).J. Conip. Physiol. 111, 309-333. Torrence, S. A. (1991).Developnun[.(Cambndge, U K ) 111, 993-1005.

LEECH NERVOUS SYSTEM DEVELOPMENT

193

Torrence, S. A., and Stuart, D. K. (1986).J. Neurosci. 6, 2736-2746. Torrence, S. A., Law, M. I., and Stuart, D. K. (1989). Dev. Biol. 136, 40-60. Tosney, K. W., and Landmesser, L. T. (1985).J. Neurosci. 5 , 2345-2358. Truman, J. W. (1984). Annu. Rev. Neurosci. 7, 171-188. Van Essen, D. C., and Jansen, J. K. S. (1977).J. Corn$. Neurol. 171,433-454. von Uexkiill, J. (1905). 2. Biol. 46 (N.S. 28); 372-402. Waddington, C. H. (1957). “The Strategy of the Genes.” Allen & Unwin, London. Wadepuhl, M. (1989).J. Exp. B i d . 143, 509-527. Wallace, B. G. (1981). In “Neurobiology of the Leech” (K. J. Muller, J. G. Nicholls, and G. S. Stent, eds.), pp. 173-195. Cold Spring Harbor Lab., Cold Spring Harbor, New York. Wallace, B. G., and Gillon, J. W. (1982).J. Neurosci. 2, 1108-1 118. Webb, R. A,, and Orchard, I. (1980). Comp. Biochem. Physiol. C 67C, 135-140. Webb, R. A,, and Orchard, I. (1981). Comp. Biochem. Physiol. C 70C, 201-207. Wedeen, C. J., and Weisblat, D. A. (1991). Development 113, (in press). Weeks, J. C. (1982a). J. Comp. Physiol. 148, 253-263. Weeks, J. C. (1982b). J . Comp. Physiol. 148, 265-279. Weisblat, D. A. (1981). I n “Neurobiology ofthe Leech (K. J. Muller, J. G. Nicholls, and G. S. Stent, eds.), pp. 173-195. Cold Spring Harbor Lab., Cold Spring Harbor, New York. Weisblat, D. A., and Astrow, S. H. (1989). Ciba Found. Symp. 144, 113-124. Weisblat, D. A,, and Blair, S. S. (1984). Dev. Biol. 101, 326-335. Weisblat, D. A., and Shankland, M. (1985). Philos. Trans. R. Soc. London, Ser. B 312, 39-56. Weisblat, D. A., Sawyer, R. T., and Stent, G. S. (1978). Science 202, 1295-1298. Weisblat, D. A., Zackson, S. L., Blair, S. S., and Young, J. D. (1980a). Science 209, 15381541. Weisblat, D. A., Harper, G., Stent, G. S., and Sawyer, R. T. (1980b). Deu. Biol. 76, 58-78. Weisblat, D. A., Kim, S. Y., and Stent, G. S. (1984). Dev. Biol. 104, 65-85. Whitman, C. 0. (1878). Q.J. Microsc. Sci. [N.S.] 18, 215-315. Whitman, C. 0. (1887).J. Morphol. 1, 105-182. Whitman, C. 0. (1892). In “Festschrift zum 70. Geburtstage R. Leuckarts,” pp. 385-395. Engelmann, Leipzig. Whittaker, J. R., Ortolani, G., and Farinelli-Feruzza, N. (1977). Dev. Biol. 55, 196-200. Willard, A. L. (1981).J. Neurosci. 1, 936-944. Wilson, E. B. (1892).J. Morphol. 6, 361-480. Wittenberg, G. (1991). Ph.D. Dissertation, Department of Biology, University of California, San Diego. Wittenberg, G., Loer, C. M., Adamo, S. A., and Kristan, W. B., Jr. (199O).J. Comp. Physiol. 167,453-459. Yau, K.-W. (1976). J. Physiol. (London) 263, 5 13-538. Young, S. R.,Dedwyler, R. D., 11, and Friesen, W. 0.(1981).J. Comp. Physiol. 141, 11 1-1 16. Zackson, S. L. (1982). Cell (Cambridge, Mass.) 31, 761-770. Zackson, S. L. (1984). Den. Bzol. 104, 143-160. Zipser, B. (1979). J. Neurophysiol. 42, 455-464. Zipser, B. (1980). Nature (London) 283, 857-858. Zipser, B., and McKay, R. (1981). Nature (London) 289, 549-554.

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GABAA RECEPTORS CONTROL THE EXCITABILITY OF NEURONAL POPULATIONS Armin Stelzer Department of Pharmacology State University of New York at Brooklyn Brooklyn, New York 1 1203

I. Introduction 11. GABAergic Inhibition: Anatomy A, GABAergic Interneurons B. Distribution of GABA Receptors C. Inhibitory Circuitry 111. GABAergic Inhibition: Physiology A. Inhibitory Postsynaptic Potentials B. Spontaneous Inhibitory Postsynaptic Potentials IV. GABAA Receptor Function: Control of the Excitability of Neuronal Populations A, Disinhibition B. Synchronization of the Inhibitory Circuit V. GABAA Receptor Function: Tetanization A, General Features B. Long-Term Potentiation C. Intracellular Regulation of GABAA Receptor Function D. Discussion VI. GABAA Receptor Function: Synchronization A. Physiological Synchronized Activity B. Disinhibition C. Synchronization of the Inhibitory Circuit D. Synchronization following Tetanization E. General Properties of Synchronization References

1. Introduction

y-Aminobutyric acid (GABA) is the major inhibitory neurotransmitter in the mammalian central nervous system (CNS). A long and often controversial discussion about the function of GABA (for review, see

Roberts, 1986) followed the first reports about its occurrence in the brain (Roberts and Frankel, 1950; Udenfriend, 1950; Awapara et al., 1950). Physiological GABA effects were first investigated at the crustacean neuromuscular junction (cf. for review, Takeuchi, 1978; Kravitz et al., 1968). 195 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 33

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Direct evidence f o r GABA as inhibitory transmitter in the mammalian CNS was established in 1967 through electrophysiological studies comparing the properties of evoked inhibitory postsynaptic potentials with responses to iontophoretically applied GABA: GABA application produces a hyperpolarization accompanied by a n increase in membrane conductance (Krnjevic and Schwartz, 1967).T h e notion that GABA is the main inhibitory transmitter in the CNS has been supported by a variety of studies that can be summarized as follows: the GABA-synthesizing enLyme glutamate decarboxylase (L-glutamate-1 -carboxylyase; EC; 4.1.1.15) is concentrated in nerve terminals of GABAergic neurons (Salganicoff and DeRobertis, 1965; Fonnum, 1968;Fonnum and Walberg, 1973;Saito et al., 1974).GABA is released from brain tissue upon $timulation by a Ca2 -dependent mechanism (Srinivasan et al., 1969; Obata and Takeda, 1969; Iversen et al., 1971; Roberts, 1974) and is eliminated from extracellular spaces by a Na+ -dependent uptake into nerve cells or glial cells (Iversen and Neal, 1968;Henn and Hamberger, 1971 ; Bloom and Iversen, 1971 ; Schrier and Thompson, 1974). There are at least two classes of GABA receptors in neurons of the CNS, termed GABA,, and GABA, receptors. GABA, receptor-mediated inhibitory synaptic potentials occur through activation of chloride conductances. Several compounds have been identified to bind specifically to the GABA,&recognition site: muscimol, isoguvacine, and 4,5,6,7tetrahydroisoxazolo[5,4-c]pyrimidin-3(2H)-one (THIP) (cf. Enna and Karbon, 1987)act as agonists and mimic physiological effects of GABA, whereas others, most notably bicuculline, block synaptic inhibition and actions of iontophoretically applied GABA in a competitive manner (Curtis and Johnston, 1970;Curtis et al., 1971). Block of postsynaptic inhibition and effects of GABA in the mammalian CNS by picrotoxin (Kellerth and Szuniski, 1966;Obata et al., 1967;Gallindo, 1969)is noncompetitive in nature (cf. Ticku, 1986).Baclofen is a specific agonist for the second major class of GABA receptors in the CNS, termed GABA, receptors (cf. Bowery, 1989).GABA binding to B receptors mediates a long-lasting hyperpolarization through G protein-mediated activation of K' conductances (-4lger and Nicoll, 1982b;Newberry and Nicoll, 1984). GABA, receptor-mediated inhibition is blocked by saclofen in a micromolar range (Bowery, 1989). GABAergic interneurons are ubiquitous and exert a powerful inhibitor) control of neuronal excitability in virtually all areas of the CNS. Under physiological conditions, the state of excitability of a cortical population represents a finely tuned balance between excitation and inhibition. Slight alterations of the efficacy of synaptic inhibition result in far greater changes in the collective behavior of neuronal populations. This +

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is particularly conspicuous in the hippocampal formation, where the activity of excitatory principal neurons-dentate granule cells and pyramidal cells-is critically controlled by intrinsic interneurons. Synaptic inhibition is mediated through both feedback (recurrent) (Spencer and Kandel, 1961; Andersen et al., 1963, 1964a,b) and feedforward (Alger and Nicoll, 1982a) circuits and the circuitry of feedforward and feedback inhibition determines the sequential activation of inhibitory pathways. GABA, receptors play a major role in the maintenance of regular cortical activity. This review focuses on the role of GABA, receptor function in the collective behavior of cortical neuronal populations, primarily in the hippocampal formation. Experimental accessibility, plasticity, and a prominent role in learning and memory account for numerous anatomical, biochemical, and physiological studies in this brain region.

II. GABAergic Inhibition: Anatomy

A. GABAERCICINTERNEURONS The anatomy of local circuit interneurons was first described in Golgi studies by Ram6n y Cajal(l893, 1911) and Lorente de N6 (1934, 1949). In the hippocampal CA1 region, at least three different populations of interneurons have been identified by these early studies: basket cells are predominantly located close to the pyramidal cell layer at the border to stratum oriens. Their axons form a dense basketlike plexus around CA1 pyramidal cell somata. Axons of interneurons at the stratum orienslalveus border arborize extensively across strata oriens, pyramidale, and radiatum (Lacaille and Schwartzkroin, 1988a). Stratum lacunosum/ moleculare interneurons arborize in strata lacunosum/moleculare, radiatum, and pyramidale with both processes, dendrites and axons. In addition, axons and dendrites can cross the hippocampal fissure to form synaptic contacts in stratum moleculare of the dentate gyrus (Lacaille and Schwartzkroin, 1988a). Anatomical, immunocytochemical, and physiological studies of hippocampal interneurons and their GABAergic nature can be summarized as follows: 1. T h e majority of interneurons in the hippocampal formation have been identified as GABAergic and inhibitory, either by GABA-antibody immunostaining (Somogyi et al., 1984; Storm-Mathisen et al., 1983;

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Gamrani et al., 1986; Sloviter and Nilaver, 1987), by glytamic acid decarboxylase (GAD)-antibody immunostaining (Frotscher et al., 1984; Ribak et al., 1978, 1981; Somogyi et al., 1983, 1984; Misgeld and Frotscher, 1986; Kunkel et al., 1986), o r by physiological criteria (Schwartzkroin and Mathers, 1978; Misgeld and Frotscher, 1986; Lacaille et al., 1987; Lacaille and Schwartzkroin, 1988a,b). 2. GABA and GAD immunostaining in the hippocampal formation is exclusively found in interneurons. Principal hippocampal (granule and pyramidal) cells are not stained. 3. GABAergic interneurons and their axonal and dendritic processes in the hippocampal formation form a dense plexus around the principal excitatory granule and pyramidal cells and arborize extensively in all hippocampal layers. GABAergic interneurons show colocalization of GABA and neuropeptides (cholecystokinin, vasoactive intestinal polypeptide, and somatostatin). In the CA 1 hippocampal subfield, somata of somatostatin-immunoreactive cells are numerous in strata oriens and pyramidale and rare in strata radiatum and lacunosum/moleculare (Kohler and ChanPalay, 1982; Morrison et al., 1982; Roberts et al., 1984; Zimmer et al., 1983; Schmechel et al., 1984; Sloviter and Nibaver, 1987). Fibers of somatostatin-positive neurons can be found in all CA1 layers (Sloviter and Nilaver, 1987). Furthermore, there is a significant overlap between GABA- and somatostatin-stained cells in CA1 (Schmechel et al., 1984; Sloviter and Nilaver, 1987). T h e distribution of somatostatin-positive cell somata in CA3 and that of other neuropeptides in the hippocampal formation is less selective (cf. Sloviter and Nilaver, 1987).

OF GABA RECEPTORS B. DISTRIBUTION

CABAergic synapses are located at both the somata and the dendrites of hippocampal pyramidal cells (Ribak et al., 1978; cf. Roberts, 1987). Inhibitory terminals are densely clustered on pyramidal cell somata and initial axon segments (Hamlyn, 1963; Ribak et ul., 1978). T h e inhibitory response of the soma (hyperpolarizing, monophasic, and chloride dependent) is primarily mediated by GABA, receptors. At least three different GABA conductances are found at pyramidal cell dendrites: a slow, G protein-mediated K + conductance and two GABA, conductances, both chloride dependent and blocked by bicuculline, but with different reversal potentials (Thalmann et al., 1981; Alger and

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Nicoll, 1979, 1982a,b; Misgeld et al., 1986). Both dendritic GABA, responses, hyperpolarizing-depolarizing at resting membrane potential, can be elicited by exogenous GABA application at the dendrites (Andersen et al., 1980a; Alger and Nicoll, 1982b; Misgeld et al., 1986). Orthodromic stimulation elicits only the hyperpolarizing GABA, component under physiological conditions. The depolarizing component can be revealed by orthodromic stimulation in the presence of pentobarbitol (Alger and Nicoll, 1982b), GABA uptake inhibitors (Hablitz and Lebeda, 1985), or 4-aminopyridine (4-AP) (Perreault and Avoli, 1988, 1989). T h e ionic nature of the underlying current is controversial, although ample experimental evidence supports the notion of a C1- iontophore coupled to the depolarizing GABAA receptor (Andersen et al., 1980a; Alger and Nicoll, 1982b; Djorup et al., 1981; Misgeld et al., 1986).

C. INHIBITORY CIRCUITRY Orthodromic stimulation of afferent fibers produces a multiphasic sequence of synaptic potentials (EPSPIfast IPSP/slow IPSP) in CA1 and CA3 hippocampal pyramidal cells and other cortical neurons. Both phases of IPSPs are activated by GABA. The fast IPSP represents the major component of both feedforward and feedback inhibition. It is blocked by bicuculline and picrotoxin and is therefore mediated by GABA, receptors (Curtis et al., 1970; Dingledine and Langmoen, 1980). T h e slow IPSP is mediated through GABA, receptors coupled via a G protein to K + channels (Alger and Nicoll, 1982b; Newberry and Nicoll, 1985; Andrade et al., 1986; Dutar and Nicoll, 1988). Antidromic stimulation of stratum oriens results in a monophasic fast IPSP mediated by somatic GABA, receptors. T h e sequential activation of inhibitory events in the hippocampal pyramidal cell is determined by both the kinetic properties of the GABA receptors involved and the circuitry of two distinct functional inhibitory pathways: a feedback (recurrent) (Spencer and Kandel, 1961; Andersen et al., 1963, 1964a,b) and a feedforward circuit (Alger and Nicoll, 1982a). Feedback inhibition in CA1 is mediated by GABAergic basket cells and the nonpyramidal class of interneurons in strata oriens/alveus (O/A) and pyramidale activated by axon collaterals of pyramidal cells (Schwartzkroin and Mathers, 1978; Schwartzkroin and Kunkel, 1985; Lacaille et al., 1987). Both basket cells and O/A interneurons receive excitatory input from CA1 pyramidal cells and in turn produce IPSPs in pyramidal cells (Lacaille et al., 1987). No experimental evidence was

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obtained for the formation of excitatory synaptic connections between pyramidal cells and lacunosum/moleculare interneurons and these interneurons may not be part of the recurrent inhibitory loop (Lacaille and Schwartzkroin, 1988b). Feedforward inhibition is activated by afferent fibers of the Schaffer commissural/collateral pathway; these fibers form direct synaptic connections with neurons of all major cell populations of the CA1 hippocampal subfield: pyramidal cells (P), basket cells (B), lacunosum/ moleculare (L/M) interneurons, and interneurons in strata orientdalveus (Lacaille and Schwartzkroin, 1988b). Electrical stimulation of this fiber pathway in stratum radiatum produces EPSPs in principal cells and interneurons of these populations (Knowles and Schwartzkroin, 1981; Buzsaki and Eidelberg, 1982; Frotscher et al., 1984; Lacaille and Schwartzkroin, 1988b). Characteristically, synaptic responses in CA 1 evoked by af‘ferent fiber stimulation in stratum radiatum are stronger in all interneuron populations with respect to pyramidal cells. In addition, the onset of the EPSPs measured in inhibitory interneurons is significantly faster than the onset in excitatory pyramidal cells (Buzsaki and Eidelberg, 1982; Lacaille and Schwartzkroin, 1988b). These characteristics of CA 1 interneurons provide the functional basis for the mediation of feedforward inhibition. Similar principles apply in CA3 (Misgeld and Frotscher, 1986) and dentate gyrus (Buzsaki and Eidelberg, 1982). Several lines of experimental evidence support the notion that L/M interneurons mediate the late (GABA,-mediated) IPSP of CA1 pyramidal cells upon orthodromic stimulation:

1. L/M interneurons are activated in a feedforward, but not feedback, manner (Lacaille and Schwartzkroin, 1988b). 2. Antidromic stimulation (activating basket cells and O/A interneurons) produces only the early GABA,-mediated IPSP. 3. T h e properties and kinetics of IPSPs evoked by L/M interneurons are similar to the late IPSP (G protein-regulated K + currents via CABA, receptors) (cf. Lacaille and Schwartzkroin, 1988b). Connections between Inhibitory Interneurons

Inhibitory synaptic potentials in inhibitory interneurons lead to disinhibitory excitation in principal cells. The main target interneurons receiving IPSPs are interneurons close to the pyramidal cell layer. L/M interneurons form synaptic contacts with the pyramidal cell layer interneurons that are coupled to pyramidal cells (Lacaille and Schwartzkroin, 1988b). Evoked inhibitory responses in L/M interneurons were observed in longitudinal slices, but not in transverse slices and not in

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paired recordings of L/M interneurons (Lacaille and Schwartzkroin, 1988a,b). O/A interneurons produce hyperpolarizing responses in interneurons in the pyramidal cell layer in a small percentage of paired recordings, but d o not seem to receive IPSPs themselves (Lacaille et al., 1987). Excitatory recurrent connections between pyramidal cells in CA 1 are rare and functionally weak compared to those of CA3 (Christian and Dudek, 1988). Disinhibitory excitation in CA 1 may functionally substitute for some of the properties attributed to recurrent excitation in other areas such as CA3. In summary, recent studies combining anatomical and physiological techniques have considerably progressed the understanding of the circuitry in CA1 and other regions of the hippocampal formation. However, the functional significance of many anatomically defined populations of neurons in the hippocampal formation (RamOn y Cajal, 1893, 1911; Lorente de NO, 1934) is not yet known. The current knowledge of the CAI circuitry can be summarized as follows (cf. Alger and Nicoll, 1982a; Buzsaki, 1984a; Lacaille and Schwartzkroin, 1988a,b): Pyramidal cells are the principal excitatory neurons and produce the fast EPSP upon stimulation of afferent fibers. Interneurons in stratum lacunosum/moleculare, basket cells in stratum pyramidale/oriens, and interneurons in alveus/stratum oriens mediate inhibitory synaptic events. All interneurons are activated in a feedforward manner following stimulation of the Schaffer collateral/commissural pathway, the interneurons in strata lacunosum/moleculare producing the late GABA,-mediated IPSP. Recurrent inhibition is mediated by basket cells and to a lesser extent by inhibitory interneurons in the strata alveus/stratum oriens region. T h e early, GABA,-mediated IPSP evoked by afferent fiber stimulation of the Schaffer collaterals represents a mixture of both feedforward and feedback activation of several populations of inhibitory interneurons. Kinetics of feedforward activation of inhibitory interneurons (feedforward inhibition) are fast with respect to those of feedforward activation of pyramidal cells (feedforward excitation) (Lacaille and Schwartzkroin, 1988b). IPSPs generated in a feedforward manner in basket cells and O/A and L/M interneurons are fast enough to count for the early IPSP following orthodromic stimulation of afferent fibers in both CA3 and CA1, although the onset of IPSPs in CAI pyramidal cells is gradual and considerably slower compared with those in CA3 (Lacaille and Schwartzkroin, 1988b; cf. Miles and Wong, 1987a, for CA3). The monophasic IPSP evoked by antidromic stimulation of fibers in strata alveus/oriens is produced by interneurons of the feedback recurrent pathway via GABA, receptors.

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111. GABAergic Inhibition: Physiology

A. INHIBITORY POSTSYNAPTIC POTENTIALS GABA-mediated inhibition occurs through two physiological mechanisms: an increase in membrane conductance and a hyperpolarization of the postsynaptic membrane. A membrane conductance increase reduces the probability of neuronal discharge by decreasing the amplitude of synchronous EPSPs. This type of inhibition has been termed “shunting inhibition? and mediates a powerful inhibitory control at membrane potentials close to the reversal potentials of ions (Ri) mediating it. The fast GABA,-mediated IPSP, through its large increase of the chloride membrane conductance, is a prime representative of this type of inhibition. T h e cessation of neuronal firing and the reduction of synchronous EPSPs are caused by the increased permeability to chloride ions, during which each depolarizing event is directly counteracted through proportionally increased chloride influx (cf. Kinjevi?, 1974). In addition, the large conductance increase reduces the electrotonic distance for concomitant EPSPs, thus confining depokarizing events and reducing the spatial summation of EPSPs and the probability of suprathreshold depolarization. With R,,, (around -75 mV) close to the resting membrane potential, the effects of GABA,-mediated IPSPs are confined to relatively small areas of the postsynaptic membrane. Such a mechanism counteracts concomitant EPSPs without affecting the general electrical activity in the rest of the neuron. Dependent upon the site of action of C1--mediated IPSPs, the effects can be very localized in individual parts of a neuronal dendritic tree, or global when generated close to strategically important sites of discharge generation (e.g., the axon hillock). Based upon theoretical calculations, “silent synapses” (R, close to K,,,,) generate the most powerful nonlinear type of inhibition, whereas excitation and inhibition interact in a linear way for synapses with R, well below R,,,, (Koch et al., 1982; Torre and Poggo, 1978): the latter type of‘ synaptic inhibition represents the second principal mechanism by which IPSPs reduce the probability of cell firing: if the IPSP has a reversal potential far below the resting membrane potential, the large hyperpolarization generated will affect large parts of the cell and reduce its general excitability. EPSPs, to reach the action potential threshold, are required to have larger amplitudes and a higher synchronization. This type of inhibition is typical for the late IPSP carried by K + currents with a low reversal potential (around -95 mV) and a relatively small increase

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of the membrane conductance. Amplitudes of synchronous EPSPs and the late IPSP sum up in a linear fashion. These functional properties of the late IPSP make this form of inhibition function as a filter for EPSPs reaching action potential threshold: EPSPs large enough to trigger cell firing remain subthreshold if the sum of the hyperpolarization plus the difference between resting membrane potential and firing threshold is larger than the EPSP. GABA,-mediated late IPSPs exhibit a slow rise time, and peak amplitudes d o not occur within 100 msec following the orthodromic stimulus: such slow kinetics are most likely due to the molecular mechanisms of G protein activation of K + conductances (Alger and Nicoll, 1982b; Newberry and Nicoll, 1984). The GABA,-mediated early IPSP exerts a profound control over both the fast EPSP and the late IPSP (Fig. 1) (Koch et al., 1982; Knowles et al., 1984; McCormick, 1989). The fast IPSP, with its short rise time and short duration, is a direct counterpart of the fast EPSP in high-frequency synaptic processing. Pharmacological blockade of GABA,-mediated inhibition results in an increase of both the amplitude and duration of the fast EPSP (Fig. 1) (cf. Knowles et al., 1984; Wigstrom and Gustafsson, 1985; McCormick, 1989). With GABA,-mediated inhibition completely blocked, a paroxysmal depolarization shift replaces the physiological EPSP-IPSP sequence. Activation of recurrent EPSPs due to blockade of GABA, receptors may to a certain extent contribute to the observed increase of the EPSP amplitude and duration, especially in brain regions with strong excitatory recurrent connections, such as the hippocampal CA3 region (Lorente de NO, 1934; Finch et al., 1983; Tamamaki et al., 1984; Miles and Wong, 1987a). But a similar potentiation of the EPSP amplitude and duration is observed in areas in which recurrent excitatory connections are absent or weak, such as the hippocampal CA1 region (Wigstrom and Gustafsson, 1985). The most probable mechanism for the observed EPSP increase is that the EPSP is directly controlled by GABA,-mediated conductances. Theoretical calculations based on time courses of the activation of fast EPSP and IPSP conductances in the circuitry of the lateral geniculate nucleus demonstrate the powerful control of the fast IPSP over the early EPSP in a locally confined area of the dendritic tree (Koch, 1985). The nonlinear strength of inhibition is a function of the delay between the activation peak of the excitatory conductance (ge) and the inhibitory conductance (gi)and is largest between 1 and 4 msec. However, steady-state (excitatory and inhibitory) inputs produce a more effective inhibitory control than do transient inputs, regardless of specific time courses of activation (Koch, 1985). The time

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FIG. 1. Transfimnation of orthodroniicallp evoked postsynaptic potentials following blockade of GABAA receptors. The top trace shows a predoniinantly GABAA-mediated response of a (:A 1 pyramidal cell upon sriniulation of the Schaffer coinmissural/collateral fiber pathway. The partial blockade of' GABA,\ receptors during the washin of the GABAA-receptor antagonist bicuculline-methiodide (50 p.M) results in an increase of the fast EPSP and the late, GAB&-mediated IPSP (middle trace). Lower trace, orthodromic stimulation of the Schaffer fiber pathway produces a suprathreshold EPSP followed by a largely increased late IPSP after the complete block of GABAA receptors (20 niin after t)icuculline bath application). The cell was held at -60 mV by depolarizing DC current injection (cell KMP: -68 mV). The action potential in the lower trace is truncated. High concentrations of divalent cations (8 n M Ca2+ and 2 inM Mg*+) were added tn the extracellular solution to prevent the generation of epileptiform responses in the presence of bicuculline.

courses for the rise and decay of transient inhibitory and excitatory conductances used in these models (which are similar to those in most central synapses) would guarantee the temporal overlap of the fast EPSPlf'ast IPSP and the resulting increase of EPSP amplitude and duration in the absence of fast inhibition under the assumption that the

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localization of excitatory and inhibitory receptors in a defined subregion of the dendritic tree is close (Koch, 1985). Blockade of the early IPSP by bicuculline leads-in addition to the increase of the duration and amplitude of the fast EPSP-also to a large increase of the late IPSP upon affernt fiber stimulation (Fig. 1) (Knowles et al., 1984; McCormick, 1989). Two possible mechanisms exist: first, disinhibition of inhibitory neurons in the absence of GABAA-mediated inhibition would functionally result in increased firing of inhibitory interneurons and an increased release of GABA. An alternative explanation is that in the absence of large GABA,-mediated conductances, K -mediated IPSPs via GABA, receptors are more effective in reaching the more negative potassium equilibrium potential (McCormick, 1989). T h e absence of GABA,-mediated shunting inhibition may also facilitate the intracellular propagation of GABA,-mediated IPSPs from distant dendritic sites of origin toward the somatic site of recording. +

B. SPONTANEOUS INHIBITORY POSTSYNAPTIC POTENTIALS In addition to generating postsynaptic potentials following stirnulation of afferent fibers, GABAA receptor-mediated chloride currents also occur spontaneously in the hippocampal formation: both spontaneous IPSPs and inhibitory postsynaptic currents (IPSCs) have been recorded in hippocampal slices (Alger and Nicoll, 1980; Collingridge et al., 1984; Miles and Wong, 1984; Gage and Robertson, 1985; King et al., 1985; Edwards and Gage, 1988; Ropert et al., 1990; Otis et al., 1990). Miniature IPSCs

Recent studies demonstrate that two types of spontaneous IPSPs/ IPSCs occur in the hippocampal formation and other brain regions: spontaneous IPSPs/IPSCs (sIPSPs, sIPSCs), evoked by spontaneous discharge of GABAergic neurons, and miniature IPSCs, which remain after cell firing and Ca2+ -dependent synaptic transmission are blocked by tetrodotoxin (TTX) and Cd2+ substitution of Ca2+, respectively (Ropert et al., 1990; Otis et al., 1990). Miniature IPSCs are significantly smaller and less frequent than discharge-evoked sIPSCs: before TTX application, sIPSCs appeared in bursts at regular intervals (every 2 sec), reaching peak frequencies of up to 200 Hz during bursts; TTX reduced the mean frequencies of sIPSCs to values between 1.2 and 1.8 Hz and the mean amplitudes to 20-28 pA (from 31-73 pH before TTX) (Ropert et al., 1990). These values were

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obtained in the CA 1 hippocampal subfield; corresponding values of miniature IPSCs recorded in the whole cell-clamp configuration in dentate granule cells were 2-20 Hz and 35 pA (Otis et al., 1990). Both miniature IPSCs and unitary sIPSCS triggered by spontaneously firing GABA cells are mediated by GABA, receptors: they are chloride dependent, blocked by bicuculline and picrotoxin, and prolonged by barbiturates (Ropert et nl., 1990). The biochemical mechanisms underlying the generation of miniature IPSPs/IPSCs are unclear: spontaneous quantal GABA release that is not dependent on discharge of GABAergic neurons has been measured in zfivo (Westerink and de Vries, 1989a). Based on observations in biochemical studies, a continuous tonic release of GABA, dependent only on the activity of the GABA-synthesizing enzyme glutamic acid decarboxylase, but not dependent on the depolarization of the presynaptic membrane, has been postulated. Independently of the GABA levels in nerve terminals, which may even be significantly elevated by blocking the GABA-degrading enzyme GABA aminotransferase, convulsions occur immediately after the activity of GAD and the rate of GABA sy9thesis is reduced to a certain extent (Tapia et al., 1966, 1975; Wood and Peesker, 1972, 1973). These observations suggest that-in addition to the conventional Ca2 - and action potential-dependent GABA reiease-newly synthesized GABA is continuously released into the synaptic cleft under physiological conditions. Although such a mechanism of GABA synthesis-release coupling could in a probabilistic manner account for spontaneous miniature events, it would hardly produce the quantal CABA release that was postulated for the occurrence of miniature IPSCs. T h e exact nature of the GABA release evoking miniature IPSCs in the absence of cell firing remains to be elucidated. Measurements of conductance changes during spontaneous IPSPsI IPSCs indicate that assuming a 16 to 20-pS main conductance state of a single GABA,, channel (Gray and Johnston, 1985; Bormann et al., 1983; Weiss et al., 1988; Edwards et al., 1989)-opening of only about 100-300 channels would be sufficient to produce the IPSP/IPSC following the discharge of a GABAergic neuron (Collingridge et al., 1984; Miles and Wong, 1984) and that only about 12-20 GABA, channels are activated during a miniature IPSC (Ropert et al., 1990). Similar estimates have been made for miniature IPSCs in dentate granule cells (Edwards Pt al., 1989).These numbers are considerably l o w e r than the number of channels activated by a single quantum of acetylcholine at the frog neuromuscular junction (1 500-2500) (Katz and Miledi, 1970; Anderson and Stevens, 1973; Pepper et ul., 1982), by a single quantum of GABA in the locust muscular junction (600-1000) (Cull-Candy, 1984), or by a +

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single quantum of glycine in the Mauthner cell (about 1000) (Faber and Korn, 1982). I n the hippocampal CAI subfield, miniature synaptic potentials are exclusively inhibitory because no miniature EPSCs could be recorded under single-electrode voltage-clamp conditions (Ropert et al., 1990). Miniature EPSPs, however, do occur in the CA3 hippocampal subfield (Brown et al., 1979) and neocortex (LoTurco et al., 1990). Miniature IPSCs, despite the activation of very few GABA channels by a single quantum of GABA, could exert a potentially powerful tonic inhibitory control of excitatory pyramidal cells. One reason is the large conductance increase by few GABA molecules at the GABAA receptor site. T h e second reason is that the amplitudes of IPSCs in CAl pyramidal cells do not correlate with their time to peak, which indicates that the electrotonic distances from the somatic site of recording are fairly homogeneous and that miniature IPSCs originate from GABAergic terminals close to the cell soma and the axon initial segment (Ropert et al., 1990). In summary, the combination of features of GABAergic inhibition, the distribution and widespread dendritic arborization of GABAergic interneurons, the ubiquitous subcellular distribution of GABAA and GABA, receptors, the concentration of GABA, receptors at strategically pivotal sites close to the cell soma, the large chloride conductance increases produced by shunting inhibition, and the generation of spontaneous IPSPs indicate a very powerful inhibitory control of a given cell population in the mammalian cortex.

IV. GABAA Receptor Function: Control of the Excitability of Neuronal Populations

Under physiological conditions, the state of excitability of a cortical neuronal population represents a finely tuned balance between excitation and inhibition. Slight alterations of the efficacy of inhibition may lead to far greater changes in the collective behavior of a population of cells. T h e prominent role of GABA,-mediated synaptic inhibition in maintaining physiological brain activity is known from early in vivo studies of GABA actions in the mammalian CNS using pharmacological blockade of GABA, receptors: block of postsynaptic inhibition and effects of GABA in the mammalian CNS by picrotoxin (Kellerth and Szumski, 1966; Obata et al., 1967; Gallindo, 1969) or bicuculline (Curtis and Johnston, 1970; Curtis et al., 1971) resulted in a generalized cellular hyperexcitability. A close correlation between reduced efficacy of

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GABAergic transmission and the generation of epileptiform activity has been demonstrated in a variety of experimental models of epilepsy (for review, see Prince, 1968; Krnjevic, 1983; Alger, 1985). In the hippocampal slice preparation, pharmacological blockade of GABAA receptors by bicuculline or penicilline (Schwartzkroin and Prince, 1980a; Dingledine and Gjerstad, 1980) or the GABAA-coupled C1- ionophore by picrotoxin (Miles and Wong, 1983; Hablitz, 1984) reliably produces enhancement of excitability and epileptiform activity in all subfields. Further support for the notion of disinhibition as a major mechanism for epileptogenesis derives from the fact that potentiation of GABA,mediated lnhibition protects against some forms of epileptiform activity, and some clinically used anticonvulsants, such as benzodiazepines and barbiturates, enhance GABA, receptor function (Nicoll et al., 1975; MacDonald and Barker, 1979). However, conclusive proof for a reduction of GABAergic inhibition as the causal mechanism in human epilepsy is elusive. Other factors that may contribute to hyperexcitable states of neuronal populations include intrinsic properties of neurons (Wong P t al., 1979; Wong and Prince, 1979; Johnston et al., 1980; Masukawa and Prince, 1984) and an increase in the excitatory drive (Ayala et al., 1973; Johnston and Brown, 1981; Wong et al., 1986).Synchronization of neuronal activity has been attributed to synaptic mechanisms [i.e., a reduction of synaptic inhibition as a major causal factor (Miles and Wong, 1986, 1987a,b; Traub et al., 1989a,b)] and nonsynaptic mechanisms, such as spread o f excitation through electrotonically coupled cells (MacVicar and Dudek, 1981; Taylor and Dudek, 1984).

,4. DISINHIBITION

Among the best-studied areas in regard to inhibition controlling the collective behavior of a neuronal population is the hippocampal CA3 subfield. T h e activity of the CA3 neuronal population represents a finely balanced activity of recurrent excitation and recurrent inhibition. Excitatory mechanisms in CA3 are potentially powerful: a single action potential in a given CA3 pyramidal cell can elicit firing in a monosynaptically coupled follower pyramidal cell (Miles and Wong, 1986). In addition, CA3 pyramidal cells are capable of firing bursts under physiological conditions (Wong and Prince, 1979). In the normal resting slice, CA3 neurons generate periodic spontaneous discharges (Wong and Prince, 1981) that occur at different frequencies in different cells and that are out of phase with one another, as shown by the lack of measur-

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able field potential and by the lack of synchronization in pairs of cells recorded simultaneously with intracellular electrodes. T h e strength of inhibitory synapses is a critical factor in controlling burst propagation between neurons: intradendritic recordings from CA1 pyramidal cells show that synaptic input in the form of CA3-generated bursts is transformed into an EPSP-IPSP sequence in CAI cells. Block of GABAA receptors results in burst discharges in CA1 dendrites (Wong and Prince, 1979; Wong et al., 1979; Traub and Wong, 1982; Masukawa and Prince, 1984). A similar control mechanism exerted by intact fast inhibition limits the propagation of bursts between CA3 pyramidal cells. Although anatomical connectivity between excitatory pyramidal cells is rather low, the few connected cells form strong excitatory polysynaptic connections between CA3 pyramidal cells. Spread of excitation is controlled by synaptic inhibition and recurrent EPSPs are not observed when inhibition is intact (MacVicar and Dudek, 1980; Miles and Wong, 1987a). Dual intracellular recording from pairs of cells show that due to the powerful inhibitory control of the recurrent pathways in CA3, burst propagation between CA3 pyramidal cells is effectively restricted (Miles and Wong, 1987a). Impairment of GABA,-mediated inhibition by picrotoxin at concentrations somewhat lower than those blocking synaptic inhibition completely leads to the activation of these polysynaptic excitatory pathways, forming connections between cells that were not connected with inhibition intact (Miles and Wong, 1987a). Unlike monosynaptic EPSPs between CA3 pyramidal cells, which can be evoked by single action potentials, it requires bursts of action potentials to elicit polysynaptic EPSPs (Miles and Wong, 1987a). Fast inhibition acts to limit the transmission of bursting. The domain of influence of a stimulated neuron progressively increases upon block of GABAA receptors. Synaptic events in a pair of neurons that are not connected with fast inhibition intact develop from nothing to a single, recurrent EPSP, then to double EPSPs, then to burst firing in the postsynaptic cell of the recorded pair concomitant with the reduction of GABA,-mediated inhibition (Miles and Wong, 1987a). Several anatomical and functional properties of the CA3 hippocampal circuitry may contribute to the powerful role inhibition exerts in the control of the network’s behavior (cf. Miles, 1990): 1. Single inhibitory connections have a conductance higher than do excitatory ones. 2. Inhibitory synapses densely cover the somata of pyramidal cells, whereas excitatory synapses terminate mainly on dendrites.

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3. Inhibitory neurons have a low firing threshold. 4. Disynaptic IPSPs have a shorter latency than do disynaptic EPSPs: fast EPSPs and the low firing threshold in inhibitory cells produce disynaptic IPSPs with latencies as short as 3-4 msec, whereas disynaptic EPSPs have typical latencies of about 12 msec (Miles and Wong, 1987a). 5. T h e connectivity for disynaptic inhibition is higher [inhibitory disynaptic interactions between pyramidal cells, 30% of cell pairs; disynaptic excitatory connections, 15% (after block of GABA, inhibition)] (Miles and Wong, 1987a). 6. There is divergence of inhibitory cell contacts (Somogyi et al., 1983; Schwartzkroin and Kunkel, 1985). 1. Derielojment of Syzchronization as a Function

of Reduced Inhibition

Properties of synchronization of cortical cell populations have been studied in a combined approach of experiments and simulation (for review, see Traub et al., 1989a,b). In an early model (Traub and Wong, 1983), several factors were proposed as requirements for synchronized bursting activity: (1) the strength of inhibitory connections must be below a critical level, (2) excitatory connections cannot be too sparse or too weak, and (3) tonic drive (to a given number of cells) produces periodic population discharges. These conditions are met in the CA2 and CA3 hippocampal subfield in which initiation of spikes (by block of GABAA inhibition) produces periodic interictal spikes in CA 1 in the intact slice, which disappear after severing the Schaffer collaterals from CA2/3 to CA1 (Schwartzkroin and Prince, 1978; Wong and Traub, 1983). Synchronized activity was generally regarded as a defining feature of epileptiform activity. However, more recent studies demonstrate that synchronized activity (to a certain extent) constitutes a form of physiological brain activity (Schwartzkroin and Haglund, 1986; cf. Traub et al., 1989a,b). In addition, simulation studies show that under physiological conditions the inhibitory control of the recurrent excitatory pathway in the CA3 hippocampal subfield may not be complete and burst firing of pyramidal cells can propagate to a certain extent within a confined area of the network (Traub e t a / . , 1989b). However, such synchronized activity under physiological conditions is small and requires the preservation of excitatory connections in a larger network: physiological synchronized activity is observed in in zriuo preparations or somewhat thicker slices (Schwartzkroin and Haglund, 1986). In the slice, both the transverse and longitudinal versions, activation of recurrent excitation and development of synchronization are closely related to synaptic disinhibition (Miles and Wong, 1987a): as GABA,mediated inhibition is reduced by picrotoxin, individual synaptic inputs

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become larger and predominantly excitatory; the interval between events and the spatial domain of spread of activity increases. Synaptically generated depolarizing potentials appear simultaneously in different cells. Synchronous EPSPs occur at a time comparable to that at which polysynaptic EPSPs are revealed following picrotoxin application. These polysynaptic EPSPs become rhythmic and their amplitude and duration grow with time of exposure to picrotoxin until the firing threshold is exceeded and synchronous discharges occur (Miles and Wong, 1987a) (Fig. 18). Synchronized bursts after complete suppression of GABA,-mediated IPSPs occur spontaneously and after stimulation. Stimulationevoked bursts require a build-up process of about 100 msec: after that delay, synchronized firing appears explosively in all cells (Traub and Wong, 1982). With inhibition completely blocked, synchronized bursting represents an all-or-none event, although the development toward fully synchronized activity is gradual (cf. Fig. 18). Furthermore, once fully synchronized bursts are developed, they can be elicited anywhere in the CA3 region (Wong and Traub, 1983). Initial stimuli leading to a synchronous population discharge are nonspecific: antidromic stimulation, potassium-induced focal depolarization of cells, synaptic input, and even activation of a single cell can initiate synchronized population discharge (Wong and Traub, 1983; Miles and Wong, 1987a). During a synchronized burst, each pyramidal cell receives a large excitatory synaptic input (Johnston and Brown, 1981; Traub and Wong, 1982). In both the model and the experiment, events triggering and associated with synchronized bursting occur in the presence of slow, GABA,mediated inhibition. Burst transmission occurs fast enough relative to the build-up of slow IPSPs, and synchronization can overturn this slow form of inhibition (Traub et al., 1987). Although there is a cascade of events, the combined expression of which leads to fully synchronized burst firing in CA3, blockade of GABA,-mediated inhibition is both necessary and sufficient to trigger large-scale synchronization processes. Fully synchronized bursting in the (complete) absence of fast, GABA, mediated inhibition in the slice represents an extreme of a range of partly synchronous population behaviors. In the experimental situation, partial synchronized neuronal activity can be obtained by several means that “tune” rather than block the efficacy of fast inhibition: application of lower concentrations of picrotoxin (or other antagonists of GABA,mediated inhibition, such as bicuculline or penicilline) results in a partial rather than a complete block of GABA,-mediated inhibition. At this stage (partial reduction of GABA,-mediated inhibition), synchronous synaptic potentials can be depolarizing, hyperpolarizing, or mixed

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(Traub et ai., 1989b). Gradual changes in the network's behavior can be studied during the wash-in process of application of GABA, antagonists (Miles and Wong, 1987a). With GABA,%-mediatedinhibition completely blocked, fully synchronized periodic bursts (with interburst intervals of several seconds) occur, affecting a large population of cells (cf. Traub et at., 1989b).

2 . Recurrvnt Excitatioii The occurrence of synchronized burst discharges put forward in the model and recorded in the CA3 hippocampal subfield (Traub and Wong, 1983) requires, in addition to a reduced inhibitory efficacy, strong functional connections between excitatory principal cells. A role for recurrent excitation for epilepsy was postulated by several authors (Burns, 1958; Ayala et al., 1973; Dichter and Spencer, 1969; Traub and Wong, 1983). Several properties of recurrent excitation contribute to the generation of synchronization. As for inhibitory neurons mediating recurrent inhibition, divergence and convergence between excitatory cells are characteristic properties of a network that exhibits recurrent excitation: a single cell can elicit polysynaptic EPSPs in more than one follower cell (Miles and Wong, 1987a). Such connectivity may represent the functional correlate of the morphological divergence of axon collaterals (Lorente de KV6, 1934; Finch et al., 1983; Tamamaki et al., 1984; Miles and Wong, 1987a). Axonal divergence facilitates the spread of activity froin one cell to a larger population during a synchronized event (Traub and Wong, 1983; Miles and Wong, 1983). The existence of several polysynaptic pathways between two cells (implying convergence of recurrent synapses) enhances the probability of recruitment of cells and the generation of synchronized bursting. In summary, experiments and simulation studies in CA3 indicate that the degree of synchronization of CA3 neurons depends on the extent to which activity can spread through recurrent excitatory pathways. Both activation of the recurrent excitatory circuitry and synchronization are contingent upon impairment of GABA,-mediated inhibition (Miles and Wong, 1987a; Traub et al., 1987, 1989a). The appearance and growth of rhythmic depolarizations in the development of synchronization reflect the growing number of cells and small groups of cells of the polysynaptic excitatory recurrent pathway firing spontaneously. The amplitude of a synchronized burst (the peak number of cells firing at once) is a nonlinear function of the strength of the fast IPSP (Traub ct al., 1987, 1989a). Synchronized bursts due to the synchronous discharge of all pyramidal and inhibitory cells represent an extreme and unphysiological

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form of population behavior and resemble the “interictal spikes”in EEG recordings of patients suffering from epileptic seizure disorders (Wong and Traub, 1983). Partial synchrony reflects subtle alterations in the balance between recurrent excitation and recurrent inhibition in the CA3 neuronal population. Rhythmical (partially) synchronized activity can be observed in many cortical areas, including the CA3 hippocampal slice, as a physiological rather than an epileptogenic condition (Schwartzkroin and Knowles, 1984; Schwartzkroin and Haglund, 1986; Schneiderman, 1986; Miles and Wong, 1987a; Traub et al., 1989b).

B. SYNCHRONIZATION OF THE INHIBITORY CIRCUIT The recruitment of latent polysynaptic excitatory connections and the development of synchronized burst activity following blockade of GABA,-mediated inhibition in the CA3 hippocampal subfield (Miles and Wong, 1987a)represent a shift (toward excitation) of a system that is well balanced by vying forces of excitation and inhibition under physiological conditions. With GABA, receptors blocked, recurrent excitatory connections become functional and consequently fully synchronized activity develops. However, simulation studies predict similar functional consequences on the collective behavior of the CA3 population if-with inhibition intact-enhanced excitatory efficacy is the experimental paradigm fed into the model (Traub et al., 1989b). In addition, a number of studies report the occurrence of highly synchronized epileptiform activity with inhibition intact or even increased (Schwartzkroinand Prince, 1980b; Gloor, 1984; Aitken, 1985; Schwartzkroin and Haglund, 1986). 1. Epileptifom Activity Evoked by 4-AP

4-Aminopyridine-evoked changes represent a prime model of epilepsy in which the development of hyperexcitability is not contingent upon a reduction of inhibitory synaptic potentials (Buckle and Haas, 1982; Rutecki et al., 1987; Perreault and Avoli, 1989; Muller and Misgeld, 1990; 1990; Aram et al., 1991). 4-AP is a very potent convulsant drug capable of inducing seizures in humans after 4-AP poisoning (Spyker et al., 1980), tonic-clonic seizures after systemic application (Pasantes-Morales and Arzate, 1981; Mihaly et ul., 1990), or electroencephalographic seizures when applied topically on the surface of the neocortex (Szente and Baranyi, 1987). A variety of different types of spontaneous activities following 4-AP application have been described in in vitro slice studies: both ictal and interictal discharges were observed in amygdala (Gean and Shinnick-Gallagher, 1989) and olfactory-cortex

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slices (Galvan et al., 1982). Interictal, PDS-like events occurred in CA3 in slices of adult animals (Matsumoto and Ajmone-Marsan, 1964; Voskuyl and Albus, 1985; Rutecki et al., 1987; Chestnut and Swann, 1988; Perreault and Avoli, 1989) and both ictal and interictal events occurred in CA3 in slices of immature animals (Chestnut and Swann, 1988). In CAI, the convulsant actions of 4-AP seem to be less effective: bursting activity was recorded in t w o studies (Voskuyl and Albus, 1985; Holsheimer and Lopes da Silva, 1989), whereas in three other studies epileptiform activity was not observed (Buckle and Haas, 1982; Perreault and Avoli, 1989; Segal, 1987). 2 . Synaptic Potentials Evoked ly 4-AP Bath application of low concentrations of 4-AP results in characteristic changes of stimulation-evoked and spontaneous synaptic potentials in the hippocampal formation and the neocortex: amplitudes of the orthodromically evoked EPSP, as well as of both phases of IPSPs in CAI pyramidal cells, are augmented (Buckle and Haas, 1982; Rutecki et al., 1987; Perreault and Avoli, 1989). Spontaneous IPSPs are greatly enhanced in amplitude and frequency following 4-AP application (Perreault and Avoli 1989; Aram et al., 1991). In addition, spontaneous EPSPs in CAI, although less prominent in amplitude and frequency than the hyperpolarizing events, are revealed by 4-AP (Perreault and Avoli, 1989). Several recent studies (Perreault and Avoli, 1989; Muller and Misgeld, 1990; Aram et al., 1991) show that 4-AP application produces rhythmic spontaneous field potentials of at least t w o different types in neocortex and hippocampus. Blockade of amino acid-mediated excitatory synaptic transmission by D-2-amino-5-phosphonovalerate(D-APV) and 6-cyano-7-nitro-quinoxaline-2,3-dione or 3-(2-carboxypiperazine-4y1)-propyl-l-phosphonate(CNQX or CPP) abolishes only type I of 4AP-induced synchronous discharges, whereas type I1 discharges persist (Perreault and Avoli, 1989; Aram et al., 1991). Type 11 field potentials last between 1 and 6 sec and occur at a frequency of 1 to 4 per minute in neocortical and hippocampal slices (Aram et al., 1991; Michelson and Wong, 1991). Intracellular correlates of 4-AP-evoked type I1 field potentials are giant and long-lasting IPSPs (6-15 mV, 1-4 sec in duration at resting membrane potential) in CAI and CA3 pyramidal cells, dentate granule cells (Perreault and Avoli, 1989; Michelson and Wong, 1991; Muller and Misgeld, 1990), and neocortex neurons (Aram et al., 1991). Although a generally accepted nomenclature of the 4-AP-evoked potentials has not yet been established, descriptive terms based on size, duration, and onset of the observed potentials allow a clear distinction from

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physiological potentials in the absence of 4-AP: giant long-lasting IPSPs, both spontaneous and stimulation evoked, are clearly distinct from physiological fast IPSPs and referred to as “slow”hyperpolarizations in some studies (cf. Perreault and Avoli, 1989). Rhythmic giant IPSPs consist of two distinct C1- components (Perreault and Avoli, 1989; Michelson and Wong, 1991) and a K + component (Segal, 1987; Muller and Misgeld, 1990). C1- and K + components of IPSPs evoked by 4-AP are, unlike stimulation-evoked IPSPs, not coupled: about 20% of giant IPSPs are either monophasic C1- or K + dependent (Miiller and Misgeld, 1990). In addition, focal application of 4-AP produced only K+-dependent IPSPs in CA3 (Segal, 1987; Muller and Misgeld, 1990), and washout of Ca2+ in the extracellular saline blocked selectively the chloride-dependent IPSP (Muller and Misgeld, 1990). Based on these observations it was proposed that giant C1- and K + IPSPs may not only be mediated by different postsynaptic GABA (GABA, and GABAB, respectively) receptors, but also by different classes of GABAergic interneurons (cf. Muller and Misgeld, 1990). In the presence of 4-AP, giant IPSPs increase in amplitude and develop into a characteristic sequence of synaptic components: between the early (fast GABA,-mediated IPSP) and a late (GABA,-mediated) hyperpolarizing component a late and long-lasting depolarizing GABA, component develops. The long-lasting depolarizing (LLD) potential consists of peak amplitudes of 2-15 mV at resting potentials and reverses between 5 and 20 mV positive of the membrane resting potential (Perreault and Avoli, 1989). I n another study, spontaneous and stimulation-evoked giant IPSPs in hippocampal principal cells have been reported to consist of three components with distinct reversal potentials corresponding to -65, -75, and -95 mV, respectively (Michelson and Wong, 1991). T h e -65- and -75-mV components represent two different GABA, responses (C1- currents) and the -95-mV component represents a GABA, response (K+ currents). The -75-mV GABA, component is identical with fast IPSPs in the absence of 4-AP (i.e., the early IPSP upon orthodromic stimulation and spontaneous IPSPs). T h e -65-mV GABA, component is most likely identical with the long-lasting depolarization that develops between the early hyperpolarizing GABA, and the late hyperpolarizing GABA, component upon 4-AP application (Perreault and Avoli, 1989). The depolarizing GABA, potential in the presence of 4-AP represents one of three components of spontaneous and orthoand antidromically evoked giant IPSPs in CA1 and CA3 pyramidal cells (Perreault and Avoli, 1989). Depolarizing GABA, responses in the presence of 4-AP are similar to those recorded in the presence of the GABA uptake inhibitor nipecotic

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acid (Hablitz and Lebeda, 1985) o r pentobarbitol (Alger and Nicoll, 1982b) or upon iontophoretic GABA application (Andersen et al., 1980a). In many of the experimental conditions under which a depolarizing GABA response is revealed, GABA concentrations at the postsynaptic site are enhanced. I t was suggested that the depolarizing component is mediated by a second class of extrasynaptic receptors that are not activated by GABA released under physiological conditions (Alger and Nicoll, 1982b). Such a mechanism could account for the long-lasting evoked and spontaneous depolarization in 4-AP because 4-AP-mediated increases of synaptic potentials are most likely due to enhancement of transmitter release, both excitatory and inhibitory (cf. Buckle and Haas, 1982; Rutecki et al., 1987; Perreault and Avoli, 1989).T h e sensitivityof GABA receptors is not enhanced in the presence of 4-AP (Perreault, 1990). Depolarizing GABA, responses in the presence of 4-AP (particularly spontaneous ones) are postsynaptic responses to bursts of GABAergic interneurons with excessive release of GABA (see below). The fact that the response to iontophoretically applied GABA is unaltered in the presence of 4-AP supports the notion of a presynaptic mechanism (Perreault, 1990). T h e depolarizing GABA, response is more sensitive to bicuculline because low concentrations of bicuculline suppress it in a selective manner (Perreault and Avoli, 1989). Based on studies performing current-source density analysis and selective ionophoretic GABA application, it is widely accepted that depolarizing GABA, receptors are mainly located in the apical dendritic tree of cortical principal cells (Andersen et al., 1980a; Thallman at al., 1981; Alger and Nicoll, 1982b; Perreault and Avoli, 1989). T h e ionic nature of the underlying current is controversial, although ample experimental evidence supports the notion of a C1- ionophore coupled to the depolarizing GABA, receptor (Andersen et al., 1980a; Djorup et al., 1981; Alger and Nicoll, 1982b; Misgeld et al., 1986; Perreault and Avoli, 1989). 3. Bursting Interrteurom

Intracellular recordings of hippocampal interneurons revealed two different responses in 4-AP in the absence of excitatory transmission: besides giant I PSPs similar to those recorded in hippocampal principal cells, large-amplitude EPSPs and bursts of action potentials were observed in other interneurons, mainly in the hilar region (Muller and Misgeld, 1990; Michelson and Wong, 1991). Paired intracellular recordings show that giant IPSPs in principal cells occur simultaneously with the bursting response in hilar interneurons (Muller and Misgeld, 1990; Michelson and Wong, 1991). Hilar interneuron bursts occur at a frequency of up to 20 per minute

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in 50 pit4 4-AP, with up to 50 action potential discharges per burst at an initial frequency of up to 300 Hz (Miiller and Misgeld, 1990). Burst responses and giant IPSPs were observed in the same interneuron, the burst response sometimes being generated during giant IPSPs (Miiller and Misgeld, 1990). Rhythmically bursting pacemaker interneurons are most likely GABAergic because synchronized discharge of these interneurons produced giant IPSPs in projected neurons (Michelson and Wong, 1991); Miiller and Misgeld, 1990). The amplitude of the depolarization wave that underlies the burst of action potentials decreased with membrane depolarization, but did not reverse due to rectification (unitary IPSPs reversed at -70 mV) (Michelson and Wong, 1991). The generation of GABA,-mediated bursts in interneurons and giant IPSPs in principal cells and interneurons is not confined to a specific hippocampal subfield, as both events persist after isolation of individual hippocampal subregions (Michelson and Wong, 1991). Several lines of experimental evidence suggest that the depolarizing (reversal at -65 mV) component of giant IPSPs (LLDs; cf. Perreault and Avoli, 1989) in principal cells and the depolarization wave that underlies bursting of interneurons-although both forms of depolarization are GABA, mediated-are different phenomena (Muller and Misgeld, 1990; Michelson and Wong, 1991). LLDs in principal cells can be reversed upon depolarization of the holding potential and do not rectify, whereas the depolarization phase underlying burst discharges in interneurons does not reverse due to rectification upon depolarization of the membrane potential (Michelson and Wong, 1991). In addition, LLDs recorded in hippocampal principal cells are usually not accompanied by action potential discharges even in cases in which the depolarization reaches firing threshold (Perreault and Avoli, 1989). This property is in marked contrast to GABA,-mediated burst responses in interneurons where bursts of action potential ride on top of the underlying depolarizing wave (Michelson and Wong, 1991). Taken together, these observations indicate that spontaneous and evoked long-lasting depolarizations in principal cells and bursts in interneurons and probably principal cells-although all mediated by GABAA receptors-are different phenomena. In summary, 4-AP reveals a variety of different responses in the neocortex and the hippocampal formation which occur in the absence of excitatory transmission and which are blocked by GABAA receptor/channel blockers bicuculline or picrotoxin (cf. Aram et al., 1991; Perreault and Avoli, 1989; Michelson and Wong, 1991; Muller and Misgeld, 1990): evoked and spontaneous giant IPSPs, including a long-

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lasting depolarizing component, type I1 (rhythmic) field responses, and depolarizing burst potentials in interneurons. In addition, the frequency of spontaneous fast IPSPs is significantly increased and spontaneous fast EPSPs that are not observed under physiological conditions occur in C A 1. Spontaneous fast EPSPs and the K component (GABA, niediated) of giant IPSPs are blocked by GABA, antagonists, most likely due to the fact that these events are driven by picrotoxin-sensitive bursts of interneurons: block of GABA, receptors abolishes the drive and the consecutive events mediated by different types of receptors (GABA, or glutamate with excitatory transmission intact). In the presence of 4-AP, hippocampal interneurons exhibit rhythmic bursts that are picrotoxin and T T X sensitive. Bursting interneurons (in the intact slice primarily in the hilus region) produce picrotoxin-sensitive giant IPSPs or depolarizing bursts in other GABAergic interneurons and synchronous giant IPSPs in principal cells in the entire hippocampal formation, including the dentate gyrus and the CA3 and CA1 hippocampal subfields. Giant IPSPs represent complex potentials consisting of at least three distinct GABA-mediated synaptic components: two GABA, components, one hyperpolarizing and one depolarizing at resting potentials, and a hyperpolarizing GABA, component. GABA release from bursting interneurons may activate the various types of GABA receptors directly or indirectly via activation of local GABAergic interneurons. Bursting of interneurons and the generation of giant IPSPs are intrinsic properties of all hippocampal subfields. These data demonstrate functional properties of GABA,, receptors evoked by 4-AP, properties that are usually attributed to excitatory transmitter-induced hyperexcitability: (1) GABAergic interneurons become synchronized in the absence of excitatory transmission; the synchronized inhibitory circuit may constitute a distinct inhibitory pathway in addition to feedback and feedforward inhibitorv pathways under 4-AP-evoked conditions; and (2) GABA acts as an excitatory transmitter producing depolarizing potentials (in principal cells) and burst responses (in interneurons). +

V. GABAA Receptor Function: Tetanization

A. GENERAL FEATURES

I. Synuptic Potentials a. Stimulation-Evoked Potentials. High-frequency (tetanic) stimulation of several afferent fiber pathways in the hippocampus induces a longterm potentiation (LTP) of synaptic transmission both in vivo and in vitro

GABA, RECEPTOR CONTROL

2 19

(Bliss and Loemo, 1973; Schwartzkroin and Wester, 1975). LTP has been proposed to serve as a cellular substrate for both learning and memory (Swanson zt al., 1982) and kindling-induced epileptogenesis (McNamara et al., 1980; Racine, 1978; Slater et al., 1985). Tetanization of afferent fibers in the CA3 region (mossy fibers in transverse slices, longitudinal association fibers in longitudinal slices) mimics the effects of pharmacological blockade of GABA, receptors on the collective behavior of the CA3 neuronal population (Miles and Wong, 1987b): after a delay of about 15 min, polysynaptic excitatory pathways are revealed. Further tetanization increases the efficacy of functional recurrent connections-both the amplitude and the duration of polysynaptic EPSPs are enhanced. Synchronous to the changes in the recurrent excitatory pathways, recurrent inhibition is reduced after tetanic stimulation; the mean amplitude of unitary IPSPs in recordings from pairs of cells is reduced from 0.7 to 0.3 mV without changing the input resistance and the IPSP reversal potential of the postsynaptic cells (Miles and Wong, 1987b).T h e spread of activity through recurrent pathways increases the synchrony of spontaneous synaptic potentials at similar time courses. Spontaneous simultaneous EPSPs emerge following the application of tetanic stimuli to both mossy fibers and longitudinal fibers. Simultaneous EPSPs grow in amplitude with time and become rhythmic similar to the development of synchronized activity during the wash-in of picrotoxin. Synchronous EPSPs following tetanization are correlated with synchronous firing in several cells as assessed by extracellular field potentials and can be evoked by activation of a single CA3 pyramidal cell (Miles and Wong, 1987b). Repeated application of high-frequency stimulus trains in CA3 stratum radiatum produces not only spontaneous and stimulation-evoked bursts of population spikes (a correlate of in vzvo interictal spikes), but also afterdischarges that resemble ictal-like, electroencephalographic seizure events (Stasheff et al., 1989). In the CA1 hippocampal subfield, repeated tetani applied to the Schaffer collateral pathway result in a similar development of highly synchronized activity (Slater et al., 1985). In intracellular recordings of CA 1 pyramidal cells, orthodromically evoked synaptic potentials, consisting of an EPSP/early IPSP/late IPSP sequence, change in a characteristic way after tetanic stimulation of stratum radiatum fibers (Fig. 2). An increase in the amplitude and duration of the EPSP is accompanied by a progressive reduction in amplitude of both phases of IPSPs, the early GABAA- and the late GABA,-mediated IPSP. T h e progressive increase in excitability produced by repetitive high-frequency stimulation is reflected in the gradual change of stimulation-evoked synaptic potentials from a multiphasic EPSP/early IPSP/late IPSP response to a monophasic depolarizing one that grows in ampli-

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CONTROL

A

POST TET.l

-Lc- LIo

-L----...........................................

POST TET.4

....................................

bid

60 MIN POST TET.6

FIG.2. (.4) Repeated tetanization results in a progressive decline of both phases of orthodroniicdly evoked IPSPs. (B) Paroxysmal depolarization shift recorded extra- and intracellularly, triggered by orthodromic stimulation 60 min after the sixth tetanus (from Stelzer r/ ti/., 1985).

tude and duration and develops into an epileptiform event similar to a paroxysmal depolarization shift (Stelzer et at., 1987) (Fig. 2). The marked alterations of synaptic potentials following tetanization are not associated with changes in the passive properties of the cell membrane such as resting potential, input resistance, inward rectification, spike accommodation, or afterhyperpolarizations (Stelzer el al., 1987) (Fig. 3). Stirnulation-evoked PSPs remain unchanged if the tetanus is applied in the presence of the N-methyl-D-aspartate (NMDA) receptor antagonist D-APV (Fig. 3). Upon washout of D-APV and further tetanization, both a reduction of orthodromically evoked IPSPs and an increase of the EPSP occur. In slices that had been tetanized in the presence of D-APV, tetanization-induced changes after washout are considerably faster compared with cells of slices that had not been previously tetanized. 0. Spontaneour Ezients. T h e occurrence of spontaneous inhibitory synaptic events in many brain regions, most notably in the hippocampal formation, has been confirmed by many studies recording depolarizing iPSPs with KC1-filled sharp electrodes or IPSCs using single-electrode voltage-clamp o r whole cell-clamp techniques (Alger and Nicoll, 1980;

CONTROL APV

POST TET.l APV

I STIM. 3.5 V

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POST TET.l WASH

POST TET.3 APV

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............................................

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FIG.3. In the presence of the NMDA receptor antagonist D-APV (10 orthodromically evoked synaptic responses remain unchanged by tetanization. Washout of APV and further tetanization produce the typical tetanization-induced changes of the EPSP-IPSP sequence. The tetanization-induced changes of synaptic potential are not accompanied by alterations of the passive cell membrane properties as assessed by membrane responses to hyper- and depolarizing current injections (from Stelzer et al., 1987).

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Miles and Wong, 1984; Collingridge et al., 1984; Gage and Robertson, 1985; King et al., 1985; Edwards and Gage, 1988; Ropert et al., 1990; Otis el al., 1990). T h e larger and more frequent events are dischargeevoked, rhythmic IPSPs triggered by spontaneously firing GABAergic interneurons (see Section 111,B). Smaller and less frequent TTXresistant miniature IPSPs constitute a second form of spontaneous inhibitory event (Ropert et al., 1990; Otis et al., 1990). In contrast to the CA3 hippocampal subfieid, no spontaneous excitatory synaptic events are measured in CAI (Brown et al., 1979; Alger and Nicoll, 1980; Ropert et al., 1990). Spontaneous EPSPs in CA1 are either absent o r too small and infrequent to be resolved with present techniques (Ropert et al., 1990). Tetanic stimulation results in characteristic changes of spontaneous IPSPs: the amplitudes and apparent frequencies of spontaneous depolarizing IPSPs measured with KC1-filled electrodes are progressively reduced to about 25% o r less of control talues after four tetani (Stelzer et al., 1987) (Fig. 4).

2 . TetarLtzation-lntluce~Modlfica tion of Receptor Sensitivity Taken together, these data indicate that the efficacy of synaptic inhibition in CA 1 pyramidal cells is progressively reduced as a consequence of tetanization. T w o main sites at which impairment of synaptic inhibition could occur are conceivable: presynaptically in GABAergic interneurons o r postsynaptically through a reduction of GABA receptor sensitivity. Exogenous GABA application, either iontophoretically or by pressure, provides a conclusive test of a possible modification of GABA receptor sensitivity. Although a detailed elucidation of the currents and receptors that mediate the exogenously evoked biphasic GABA response CON

CON

25C

FIG.4. Amplitudes of spontaneous inhibitory postsynaptic potentials per minute are progressively reduced by tetanization (from Stelzer el al., 1987). CON, control; PT, post tetani.

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(Andersen et al., 1980a) is not yet available, it may be inferred from the features of orthodromically evoked IPSPs and the subcellular distribution of GABA receptors that the biphasic, hyperpolarizing-depolarizing GABA response following iontophoretic application is mediated by both types of GABA, receptors (dendritic and somatic, depolarizing and hyperpolarizing, both coupled to chloride ionophores). A possible contribution of GABAB receptors has not been demonstrated, but based upon the kinetics of GABA,-mediated potassium currents and the distribution of GABA, receptors, it can be assumed that GABA, receptors are activated under conditions that produce a biphasic GABA response: the depolarizing (GABAA) response demonstrates that iontophoretically released GABA reaches sufficient concentrations at dendritic sites of the cell at which GABA, receptors are presumably located. T h e biphasic hyperpolarizing-depolarizing response upon iontophoretic application of GABA in CAI pyramidal cells (Andersen et al., 1980a) decreases progressively following application of tetanic stimuli: the impairment of both phases of the GABA response together with orthodromically evoked IPSPs (Fig. 5) suggests that the loss of synaptic inhibition is due to a reduction of the sensitivity of the GABA receptors mediating the iontophoretic response, i.e., GABA, and GABA, receptors. T h e NMDA receptor antagonist D-APV prevents all changes observed after tetanization of afferent fibers: (excitatory and inhibitory) synaptic potentials and responses to iontophoretically applied GABA remain unaltered by tetanization in the presence of NMDA antagonists. Changes of the collective behavior of the CA1 population-the development of epileptiform and synchronized activity-are also prevented by sufficiently high doses of D-APV (>5 pM) added to the extracellular solution. These data indicate that tetanization effects in the CAI hippocampal subfield are contingent upon activation of NMDA receptors: washout of D-APV and further tetani produce the usually observed tetanization-induced changes of synaptic potentials and receptor sensitivity, although the progression of alterations after washout of D-APV and further tetanization is somewhat faster compared with slices that had not been exposed to tetani in the presence of APV. The notion that the changes of synaptic inhibition occur through NMDA receptor action is further supported by the observation that the NMDA receptor sensitivity-as assessed by various parameters (rise time, amplitude, and duration of the response to iontophoretically applied NMDA)-is greatly enhanced by tetanization (cf. Fig. 5 ) . These data indicate that the efficacy of GABAergic inhibition is impaired postsynaptically through a reduction of GABA receptor sen-

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F i c . 3. Tetanic stimulation changes GABA and NMDA receptor sensitivity. (A) The biphasic hyperpolarizing-depolarizing responses to iontophoretically applied GABA as well as both IPSP components decline following tetanization. The responses to iontophoretically applied NMDA increase in amplitude and duration following tetanization.

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sitivity. To address the question of possible presynaptic modifications of GABAergic transmission, a series of experiments was performed to study effects of tetanization in interneurons located in or close to the CA 1 pyramidal cell layer. These interneurons, most likely inhibitory basket cells, have been characterized by electrophysiological and anatomical experiments (cf. Schwartzkroin and Mathers, 1978). Cells were chosen based upon membrane responses to depolarizing and hyperpolarizing current injection (cf. Fig. 6, control inset): brief spikes, short time constants, small input resistance, lack of inward rectification during hyperpolarizing current pulses, large afterhyperpolarization. The late IPSP upon orthodromic stimulation of the Schaffer collaterals was less or was not expressed in interneuron recordings (cf. Figs. 6 and 7). Tetanization produced similar changes of the orthodromic EPSPIPSP sequence as observed in CAl pyramidal cells following tetanization of the Schaffer collaterals (Figs. 6 and 7). The reduction of the IPSP and increase of the EPSP in interneuron cells occurred during the same time course as in CA1 pyramidal cells. Iontophoretic GABA application produced a biphasic hyperpolarizing-depolarizing response similar to the responses in CA1 pyramidal cells. In nonpyramidal cells, however, tetanic stimulation leads to an increase of the depolarizing iontophoretic GABA component (Fig. 6). The increase of the depolarizing response to iontophoretic GABA application resulted in burstlike events as soon as post tetanum one (Fig. 6). The orthodromically evoked IPSP following tetanization was reduced but remained hyperpolarizing after the first and second tetanus and did not match the large depolarizing response to exogenously applied GABA. The depolarizing GABA response in interneurons was accompanied by a large increase in action potential firing (Fig. 6). In this respect, depolarizing GABA actions resemble the burst responses in interneurons in the presence of 4-AP (cf. Muller and Misgeld, 1990; cf. Section IV,B). Depolarizing spontaneous fast IPSPs in interneurons are similarly reduced following tetanization (Fig. 7). However, large spontaneous depolarizing potentials develop, usually between the third and fourth tetani (Fig. 7). Spontaneous depolarizing potentials grow in amplitude upon further tetanization and eventually develop into large PDS-like events with bursts of action potentials riding on them (Fig. 7). At this stage of excitability, orthodromic stimulation (stratum radiatum fibers) (B) In the presence of D-APV,tetanization does not produce alterations of G A B A responses or orthodromically evoked synaptic potentials. Washout of APV and further tetanization result in changes of G A B A receptor sensitivity and inhibitory synaptic potentials commonly observed after tetanization (from Stelzer et al., 1987).

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POST El.1

POST TET.2

' !

400ns

'

,

GABA

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H

10 nV

FIG. 6. Tetanization of stratum radiatum fibers increases the depolarizing component of the biphasic hyperpolarizing-depolarizing control response to iontophoretically applied GABA in nonpyramidal cells. Orthodromically evoked synaptic potentials (upper row) are measured at the cell's resting potential (-67 mV); recordings of GABA responses were performed at the more depolarized potential of -61 mV (obtained by depolarizing DC current injection). The inset (upper row,left) shows the cell's IV characteristics, which are profoundly different from those of CAI pyramidal cells (cf. Fig. 3). The growth of the depolarizing GABA response was accompanied by high-frequency action potential discharges (bottom, post tet. 1 and 2) (note: frequency of discharges supersedes the chart's ability to monitor full lengths of action potentials).

produced burstlike depolarizing potentials in interneurons (Fig. 7, top). Spontaneous giant depolarizations and bursts occur infrequently, are not rhythmic, and neither the frequency nor the probability of occurrence is dependent on the membrane potential. Several lines of experimental evidence indicate that the depolarizing giant events are GABA, receptor mediated, despite the fact that amplitude and frequency of small spontaneous depolarizing IPSPs progressively diminish-most likely due to a loss of GABA, receptor sensitivity (see below). Giant potentials are not observed when bicuculline is applied at these later stages of tetanization. In addition, giant depolarizing potentials occur in slices in which the CA2/CA3 pacemaker region for synchronized activity in CA 1 is removed by dissection. Excitatory synchronized activity, including epileptiform activity, is not spontaneously generated in CA 1 (Brown et al., 1979; Alger and Nicoll, 1980; Ropert et at., 1990) due to the specific anatomical and functional properties of the circuitry, in particular the lack of recurrent excitatory connections in this area (Wong

227

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POST TET.3

POST TET.4

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SPONTANEOUS

J J -+ % & +

*

FIG. 7. Spontaneous giant IPSPs and bursts in nonpyramidal cells. Repeated tetanization in a nonpyramidal CAI neuron (IV characteristics as in cell of Fig. 6 are not depicted) produces spontaneous giant depolarizing potentials and bursts, which can be blocked by GABAA antagonists (not shown). Giant IPSPs and bursts occur despite the progressive fading of the amplitude and frequency of fast, spontaneous control IPSPs. The upper row depicts stimulation-evoked potentials, which resemble bursts at increased stages of excitability following the third and fourth tetanus.

and Traub, 1983; Schwartzkroin and Prince, 1978). The question as to whether giant IPSPs and GABAergic bursts occur as highly synchronized discharges of a large number of cells in the presence of the remainder of (reduced) GABA, sensitivity (after three or four tetani) or whether the synchronized GABAergic events are mediated by a different kind of receptor, the sensitivity of which is actually enhanced following tetanization (cf. Fig. 6), remains to be elucidated. The fact that tetanization-induced spontaneous giant IPSPs occur exclusively in nonpyramidal neurons in which the depolarizing iontophoretic GABA response is actually enhanced following tetanization (Fig. 6) strongly favors the latter notion. The infrequency of giant IPSPs after tetanization may be due to a lesser expression of synchronization of inhibitory interneurons compared with that in 4-AP. Another factor may be the isolation of the CA 1 hippocampal subfield in which inhibitory bursting activity can be generated (as inferred from 4-AP experiments; cf. Section IV,B), but to a lesser extent compared with other hippocampal

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subfields, in particular the hilar region (Michelson and Wong, 1991; Muller and Misgeld, 1990). I n summary, tetanic stimulation of Schaffer collaterals results in the following changes in CA 1-located interneurons: 1 . Orthodromically evoked PSPs shift toward increases in the EPSP and a reduction in IPSPs similar to CA1 pyramidal cells. 2. T h e depolarizing component of the biphasic hyperpolarizingdepolarizing control response to exogenous GABA is largely increased and the hyperpolarizing component is decreased. A functional evaluation of tetanization-induced changes in interneurons has to take into consideration the complex circuitry and connectivity of interneurons. However, the increased excitability of feedforward interneurons following tetanization of afferent pathways (Buzsaki and Eidelberg, 1982) (cf. Figs. 6 and 7) is indicative of an increase of inhibitory efficacy at presynaptic sites (with respect to CA 1 pyramidal cells). With respect to interneurons, however, underlying mechanisms of tetanization-induced interneuron hyperexcitability-in analogy to CA 1 principal cells-are most likely postsynaptic. Taken together, the changes produced by tetanization of afferents on the pre- and postsynaptic inhibitory efficacy in the hippocampal CA1 region may be opposite. Mechanisms as t o how alterations of the inhibitory circuit may result in a net increase of excitability are discussed below (cf. Sections V,B and V,D). SynchroniLed giant IPSPs and burst potentials, after repeated tetanization (Fig. 7), are reminiscent of 4-AP-induced changes of the inhibitory circuitry. Underlying mechanisms remain to be elucidated. T h e tetanization-induced shift of interneuron GABA sensitivity toward depolarization and a possible role of GABA acting as excitatory transmitter may, however, play a major role in the generation of synchronized inhibitory bursts (cf. Sections IV and VI). Another property of CA1 neurons, which may play a role in cellular synchronization, is characteristically altered following Schaf€er collateral tetanization: rhythmical membrane potential oscillations (MPOs) (Fujita and Sato, 1964) occur in hippocarnpal neurons in CA1 and CA3 (Leung and Yim, 1988). MPOs are subthreshold oscillations of the membrane potential of hippocampal neurons at frequencies between 3 and 11 Hz (Leung and Yim, 1988). Each cycle of an MPO probably consists of a depolarizing phase caused by N a + and Ca'+ currents that are below the spiking threshold, and a repolariation phase by K + currents (Leung and Yim, 1988). Spikes and hippocampal membrane oscillations are not linked, although membrane oscillations occur at membrane potentials close to the spiking threshold

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and most oscillations are accompanied by action potential firing. MPOs can be triggered by depolarizing current injection (Leung and Yim, 1988). The threshold of membrane potential oscillations is significantly reduced after tetanic stimulation (Fig. 8, bottom right). The lowering of the threshold for MPOs is a progressive development following tetanization: after four or more tetani, some cells exhibited large MPOs close to the resting potential. The frequency of spontaneous discharges in cells was largely increased despite the fact that the firing threshold remained unaltered following tetanization. MPOs occur in principal cells and interneurons under control conditions. The smaller gap between resting potential and action potential threshold in interneurons may have a more significant impact for interneuron oscillations after tetanization. Block of synaptic transmission by omission of Ca*+ attenuates MPOs (Leung and Yim, 1988). MPOs are sensitive to GABA shunting and are

CONTROL

STIM.

L POST TET.l

400 ms

FIG.8. Threshold for membrane potential oscillations (MPOs) is lowered by tetanization. Orthodromically evoked synaptic potentials (top row) are recorded at resting membrane potential (-73 mV); GABA responses are recorded at a depolarized membrane potential of -68 mV (by DC current injection). Before tetanization, the MPO threshold was about -63 mV (not shown). Action potential threshold of about -60 mV remained unchanged by tetanization. The cell's IV characteristics are typical of a CAI pyramidal cell. Similar observations are made in nonpyramidal interneurons. During the responses to iontophoretically applied GABA, MPOs were completely abolished (recording at bottom, right).

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completely abolished during the biphasic response to iontophoretically applied GABA (Fig. 8). In summary, two spontaneous events in CAI interneurons are more pronounced after tetanization of stratum radiatum fibers. T h e differentiating features of depolarizing (giant) IPSPs and MPOs are severalfold. T h e major difference lies in the fact that MPOs occur in principal cells and interneurons whereas giant IPSPs occur exclusively in cells with interneuron properties. T h e regularity and frequency of MPOs increase with the level of depolarization. Giant IPSPs (in CA 1) are solitary, infrequent events, the frequency of which is not dependent on membrane potential changes. Tetanization lowers the threshold for the generation of MPOs that are observed in controls at membrane potentials close to the firing threshold and are reliably triggered by depolarizing current injection. Giant IPSPs are not observed until several tetani are administered (usually after three to four tetani). Synaptic currents (GABA-mediated chloride currents) drive giant IPSPs and intrinsic currents (Na+, Ca2+, K + ) drive MPOs, but blockade of synaptic transmission attenuates the occurrence of MPOs (Leung and Yim, 1988). 3. Time Course of Effects of Tetanimtion

The tetanus-induced changes in excitability discussed herein, including the reduction of the GABA,-mediated early IPSP and the sensitivity of GABA,, and NMDA receptors, usually develop around I5 min after the application of a given tetanus and are fully expressed after about 30 min following individual tetani: if no further tetani are applied, the tetanus-induced effects are long-lasting and remain until the end of the experinient-up to 4 hr as assessed intracellular and u p to 8 hr measured extracellularly. Further tetanization (in 30-min intervals) resulted in a progressive development of the changes described. T h e time course and duration of the tetanization-induced effects indicate the involvement of intracellular regulation processes [second-messenger-mediated phosphoryiation processes o r activation of early genes (cf. Cole et al., 1989)]. Tetanization evokes several short-term changes (within 15 min following the tetanus) that affect GABA, and GABA, receptor sensitivity. Shifts of the ionic equilibria of ions mediating GABAergic inhibition (Cl- and K + ) contribute to these short-term changes, which reverse within a few minutes (Wong and Watkins, 1982; Thompson and Gahwiler, 1989a,b,c).N o change of the ionic equilibria and reversal potential of either phase of the orthodromic IPSPs is observed 20 min after individual high-frequency trains (Stelzer el al., 1987).

GABAA RECEPTOR C O N T R O L

23 1

B. LONG-TERM POTENTIATION Learning and memory processes are likely to involve long-lasting, use-dependent alterations in the efficiency of synaptic communication. Such alterations may result from changes at existing synapses (Hebb, 1949; Eccles, 1953) or from alterations in the number of functional synaptic connections (Tsukahara, 1981). Groups of synaptically associated cortical cells might represent a basis for information storage and might be formed if collateral synapses between them are strengthened (Lorente de NO, 1949; Hebb, 1949; Marr, 1971; Gardner-Medvin, 1976; Abeles, 1982; Miles and Wong, 1987b). A valuable model for investigating long-lasting synaptic alterations is long-term potentiation. LTP can reliably be induced by a short, highfrequency stimulus to afferent pathways in many areas of the mammalian CNS, most notably in the hippocampus (Bliss and Loemo, 1973). Several features of LTP support the notion of LTP as a possible physiological substrate for learning and memory: its long-lasting duration (up to days and weeks following tetanization in vivo) (Bliss and Loemo, 1973) and its relative input specificity (Dunwiddie and Lynch, 1978; Andersen et al., 1980b), cooperativity (McNaughton et al., 1978; Lee, 1983), and associativity (Levy and Steward, 1979; Sastry et al., 1986). A number of reviews cover various aspects of LTP (cf. Collingridge and Bliss, 1987; Malenka et al., 1989; Kennedy, 1989). The role of synaptic inhibition in LTP, in particular the role of GABA, receptors, will be the focus of the following discussion. Synaptic inhibition has been ruled out as a major factor in the expression of LTP by earlier reports (Haas and Rose, 1982; Wigstrom and Gustafsson, 1985; Abraham et al., 1987; Taube and Schwartzkroin, 1987). This conclusion may be premature in the light of the marked reduction of the inhibitory tone following high-frequency stimulation of afferent fibers and the prominent role of the inhibitory circuit in the control of the excitability of hippocampal population activity. 1. Physiological Stimulus LTP can reliably be obtained by a standard tetanus (duration between 200 psec and about 2 sec, frequencies between 50 and 100 Hz). However, a wide variety of stimuli of afferent fiber pathways produce LTP-like synaptic plasticity (cf. Gustafsson and Wigstrom, 1988). The survey of the different stimulation patterns that are capable of inducing LTP leads to the conclusion that depolarization of the postsynaptic membrane to a certain degree represents a necessary requirement for

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the induction of LTP (Douglas et al., 1982; Wigstrom and Gustafsson, 1985; Sastry et al., 1986). Recent studies have examined stimulation patterns similar to those produced in vivo. Short, high-frequency bursts (four-pulse bursts, 100 Hz) applied to the Schaffer commissural projections to the CA1 subfield elicited LTP in a repetition window between 0.1 and 2 sec: burst intervals of 2 sec and longer were not very effective whereas 200-msec repetition intervals were most effective in eliciting LTP (Larson et al., 1986). ‘These stimulation patterns resemble spike discharge patterns of hippocampal neurons in animals exposed to learning situations. Moreover, when bursts were applied to two different dendritic inputs to CAI pyramidal cells, the second bursts elicited robust LTP at an appropriate interval (200 msec) even when the t w o input sites innervated completely different regions of the postsynaptic cells (Larson and Lynch, 1986). LTP did not occur when the inputs were stimulated simultaneously o r when the second burst was delayed by 2 sec (Larson and Lynch, 1986). The data indicate that a single burst produces a transient, spatially diffuse priming effect that modifies subsequent bursts to produce synaptic plasticity in a spatially confined area. Similar stimulation and timing patterns (bursts consisting of about four spikes) occur physiologically in the hippocam pal formation: CA3 pyramidal cells generate periodic, spontaneous burst discharges (Wong and Prince, 1979) very similar to those applied in the patterned stimulation experiments by Larson et al. (1986). As discussed in Section VI,A (cf. Traub et al., 1989b), partially synchronized burst firing constitutes physiological brain activity, both in uiuo and in vitro. The functional state of GABA, receptors is a critical factor in the generation of synchronized activity (cf. Miles and Wong, 1987a; cf. Fig. 18) and determines the number of axonal fibers synchronously activated. Taken together, it is conceivable that synchronous burst discharges in the disinhibited CA3 hippocampal subfield may provide the cooperative activation of patterned afferent fiber stimulation required to elicit longlasting synaptic plasticity in CAI. Furthermore, the similarity of the temporal parameters of the priming effect [maximal LTP effect at frequencies of about 5 Hz (Larson and Lynch, 1986)l and the theta rhythm (4-7 Hz) that occurs in the hippocampus during learning episodes supports the notion of such periodic, synchronous bursting as a correlate of the LTP-inducing input. T h e output pattern controlled by the state of synchronized activity of the (entire) CA3 population or specific clusters of CA3 cells may represent a crucial factor in generating the physiological stimulus producing synaptic plasticity effects in CA1.

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2. Induction of LTP Induction of LTP in the CA1 hippocampal subfield is contingent upon the activation of NMDA receptors (Collinridge et al., 1983; Harris et al., 1984; Wigstrom and Gustafsson, 1985). NMDA receptor activation requires a sufficient depolarization of the cell membrane (Douglas et al., 1982; Wigstrom and Gustafsson, 1985; Malinow and Miller, 1986; Sastry et al., 1986) to alleviate the Mg2 block of NMDA channels (Mayer et al., 1984). This notion is supported by intracellular studies demonstrating that the EPSPs of single pulses or weak tetanic stimuli that are not sufficient to trigger LTP alone produce LTP when combined with intracellular depolarizing current application (Kelso et al., 1986; Sastry et al., 1986; Wigstrom et al., 1986). Tetanization fails to elicit LTP when the neuron is kept hyperpolarized (Malinow and Miller, 1986). A reduction of the inhibitory input during the depolarizing, LTP-inducing stimulation phase greatly facilitates the induction of LTP in both hippocampus (Wigstrom and Gustafsson, 1983) and neocortex (Artola and Singer, 1987). In addition, properly timed inhibitory inputs reduce or prevent the occurrence of LTP (Douglas et al., 1982). In neocortex, the block of GABAA-mediated inhibition represents a necessary condition for the induction of LTP by tetanic stimulation (Artola and Singer, 1987). These findings suggest that GABA, receptors in the feedforward circuitry control the amount of postsynaptic depolarization necessary for the activation of NMDA receptors in the induction phase of LTP. Two possible mechanisms may contribute to the disinhibition of the feedforward inhibitory pathway: first, a decrease in the efficacy of synaptic inhibition (either pre- or postsynaptic), and second, functional synaptic connections between inhibitory interneurons (Misgeld and Frotscher, 1986; Lacaille et al., 1987; Freund and Antal, 1988; Lacaille and Schwartzkroin, 1988b). +

3. Maintenance oJ: LTP LTP is commonly measured by extracellular or intracellular recordings. In extracellular recordings in the pyramidal cell layer, orthodromic stimulation evokes a characteristic sequence of field potentials consisting of an early antidromic spike (more pronounced in in vivo recordings) and a later negative-going orthodromic population spike, superimposed upon a positive synaptic wave, representing dendritic excitatory postsynaptic potentials. High-frequency stimulation increases largely the negative-going orthodromic population spike and the positive-going EPSP, leaving the negative-going antidromic spike almost unchanged

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(Bliss and Loemo, 1973). T h e population spike amplitude reflects the number of cells firing upon orthodromic conditioning stimulation. Dendritic excitatory potentials can be more directly assessed through extracellular recordings of field potentials in the dendritic area (usually recorded in stratum radiatum of CA1 o r CA3). In intracellular recordings, orthodromic stimulation of afferent fibers evokes a sequence of EPSPsIPSPs in pyramidal cell somata (cf. Section 111,A). T h e most common parameter measured in LTP experiments is the increase in the EPSP amplitude (with fast inhibition often blocked) or the EPSP slope. Recently, intradendritic (Taube and Schwartzkroin, 1988) and whole cell patch recordings in the slice (cf. Malinow and Tsien, 1990) have been performed to elucidate LTP mechanisms. Most studies focus on modifications of excitatory amino acid-mediated transmission properties underlying the expression of long-term potentiation: persistent modifications of synaptic transmission may occur presynaptically (Dolphin et al., 1982; Bekkers and Stevens, 1990; Malinow and Tsien, 1990) or postsynaptically. Possible postsynaptic modification sites following tetanization include a-amino-3-hydroxy-5methyl-4-isoxazole propionic acid (AMPA) receptors, which mediate fast EPSPs (Muller et al., 1988; Kauer et al., 1988; cf. Kennedy, 1989), and NMDA receptors, which mediate slow components of excitatory postsynaptic potentials. Activation of NMDA receptors is a prerequisite (at least in some areas, such as the hippocampal CAI subfield) for the induction of LTP (Collingridge et al., 1983). A recent study indicates that NMDA receptor-mediated postsynaptic potentials may also be persistently enhanced following tetanization (Bashir et al., 1991). a. IPSP Reductionfollowing Tetanization. Following tetanic stimulation of afferent fibers, orthodromically evoked IPSPs were found to be unchanged, increased, or decreased using the same stimulation protocol (Yamamoto and Chujo, 1978; Misgeld et al., 1979; Abraham et al., 1987; Haas and Rose, 1982; Taube and Schwartzkroin, 1988). A typical sample of evoked IPSP alterations following tetanization was reported by Abraham et al. (1987), who found IPSP increases in 8 of 19 CA1 pyramidal cells, a decrease in 3 cells, and no IPSP changes in 8 cells. Similar inconsistent changes of evoked IPSPs were also obtained in intradendritic recordings (Taube and Schwartzkroin, 1988). A decrease of stimulationevoked IPSPs after tetanic stimulation was measured in CA3 (Yamamoto and Chujo, 1978; Misgeld et al., 1979; Misgeld and Klee, 1984; Miles and Wong, 1987b) and CA1 pyramidal cells (Haas and Rose, 1982; Larson and Lynch, 1986, Abraham et al., 1987; Stelzer et al., 1987; Taube and Schwartzkroin, 1987, 1988). IPSP reductions following tetanization were most commonly observed following repeated long and strong tetaniza-

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235

tions (Abraham et al., 1987; Misgeld and Klee, 1984). However, marked reductions of orthodromically evoked IPSPs were observed following prime pulses, which are relatively weak stimuli but resemble in viuo stimulation patterns (Larson and Lynch, 1976). IPSP increases following tetanization in CA3 were found to be pathway specific; decreases were consistently heterosynaptic in nature (Misgeld et al., 1979). Haas and Rose (1982) attributed the observed decrease in the stimulation-evoked IPSPs to the fact that the increase in the fast EPSP produced the concomitant reduction of the overlapping early IPSP. T h e conclusion of most studies was that tetanization-induced changes of IPSPs are not significant and inhibition does not play a major role in the maintenance of LTP (cf. Haas and Rose, 1982; Taube and Schwartzkroin, 1987; Abraham et al., 1987). As discussed in Section V,A, tetanization produces opposite pre- and postsynaptic changes of the inhibitory efficacy: an increase in interneuron excitability and a reduction of postsynaptic GABA sensitivity. Opposite pre- and postsynaptic changes of the inhibitory efficacy could account for inconsistent changes of orthodromically evoked IPSPs in LTP studies. If so, to what extent and under what conditions are preand postsynaptic alterations of inhibitory synaptic efficacy expressed after tetanization? b. GABA Sensitivity. As discussed in Section V,A, GABA receptor sensitivity in CA 1 is characteristically altered after tetanization of the Schaffer collaterals: in CAI pyramidal cells, both the hyper- and depolarizing components of the response to iontophoretically applied GABA are progressively reduced after application of repeated tetani. T h e control GABA response is most likely composed of several GABA receptor components, including GABA, and GABA, components. T h e impairment of the GABA response together with both phases of the orthodromically evoked IPSPs (Figs. 5 and 8) indicates that the loss of synaptic inhibition is due to a reduction of the sensitivity of GABA receptors, both GABA, and GABA,. I n nonpyramidal cells with characteristics similar to those recorded in CAI basket cells (Schwartzkroin and Mathers, 1978), iontophoretic GABA application produced a biphasic hyperpolarizing-depolarizing response similar to the responses in CA1 pyramidal cells. In these cells, however, tetanic stimulation produced an increase of the depolarizing iontophoretic GABA component (Fig. 6).The reduction of IPSPs in these cells occurred at the same rate as in other neurons in which the biphasic GABA response progressively faded. The increase of the depolarizing iontophoretic GABA response, however, occurred somewhat faster and iontophoretic GABA application after the second tetanus produced

burstlike events that outlasted the duration of GABA application considerably (Fig. 6). These data demonstrate specific alterations of the GABA sensitivity in different cell types of the CAI area following tetanization. In a previous study, GABA sensitivity was reported to be unaltered in slices following LTP (Scharfman and Sarvey, 1985). A possible explanation for these data may be found in opposite alterations of GABA sensitivity in CA 1 cells. In pyramidal cells, which outnumber interneurons t o a large extent (Cassell and Brown, 1977), both components of the hyperpolarizing-depolarizing control responses are progressively reduced following tetanization. In at least one class of nonpyramidal cells located close to the pyramidal cell layer, the depolarizing GABA response is highly increased. Extracellular measurements of GABA responses may not reveal such specific alterations and may not be the adequate approach in the light of opposite changes of GABA sensitivity following tetanization. c. Interneuron LTP. T h e results of recordings from interneurons demonstrate increased basket cell activity following tetanic stimulation of afferent fibers: LTP in CA1 interneurons was reported in in vivo (Buzsaki and Eidelberg, 1982; Kairiss et al., 1987) and in aztro (Taube and Schwartzkroin, 1987; Abraham et al., 1987) studies. The degree of interneuron LTP seems to be higher in in TWO preparations than in the slice (Abraham et al., 1987). It was hypothesized that increases in excitability following tetanic stimulation of afferent fibers occur when the degree of potentiation of the feedforward inhibitory system is smaller than the increase of excitatory transmission properties (Wilson et al., 1981; Abraham et al., 1987). Although the exact degree of tetanization-induced potentiation of basket cell activity remains to be determined, it is widely accepted that basket cell activity (in CA1) is not decreased following tetatiizatiori and that the potentiation of excitability following high-frequency stimulation cannot be explained by the observed changes in interneuron activity (Taube and Schwartzkroin, 1988). Additional factors that may contribute to a presynaptic strengthening of the inhibitory system include the activation of an increased number of inhibitory interneurons after the tetanus. Enhanced efficacy of inhibition may he further achieved by the synchronization of the inhibitory circuit (Fig. 7) and by the enhanced sensitivity of the depolarizing GABA response in interneurons (Fig. 6). In summary, tetanization results in increased interneuron activity (Buzsaki and Eidelberg, 1982; Kairiss et al., 1987; Taube and Schwartzkroin, 1987; Abraham et al., 1987) and an increased number of recurrent interneurons activated by the control stimulus. The observation of orthodromically evoked “unchanged” IPSPs measured in somatic intracellular recordings after a single tetanus implies, therefore, an actual reduction of the postsynaptic GABAergic

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efficacy in the presence of increased interneuron excitability. Mechanisms that strengthen synaptic inhibition presynaptically (activation of a larger number of interneurons, interneuron LTP) are likely pathway specific and may mask the actual reduction of the reduced postsynaptic GABA sensitivity. Repeated or stronger high-frequency stimulation may result in an overall reduction of the inhibitory efficacy by reducing (postsynaptic)GABA sensitivity to a larger degree than enhancing (presynaptic) interneuron activity. At this stage of excitability, orthodromically evoked IPSPs are generally reduced (for a more elaborate discussion, see Section V,D). d. E-S Potentiation. A number of studies demonstrate that potentiation of somatic excitability (measured by the population spike amplitude) exceeds by far the increase in the population EPSP (reflecting excitation at the dendritic site). This enhanced EPSP-spike relationship, termed E-S potentiation (Andersen et al., 1980b),has been observed in both dentate gyrus (Bliss and Loemo, 1973; Wilson etal., 1981)and CA1 (Andersen et al., 1980b; Reymann et al., 1987;Abraham et al., 1987; Taube and Schwartzkroin, 1987, 1988). Intradendritic recordings in CAI pyramidal cells confirm the poor correlation between dendritic and somatic LTP: population spike amplitudes were often potentiated without concomitant changes in the intradendritically recorded EPSP (Taube and Schwarukroin, 1988). A dendritic EPSP increase is always accompanied by population EPSP increases and vice versa (Taube and Schwartzkroin, 1988). These data demonstrate that alterations in the EPSP (measured intra- or extracellular) are not a prerequisite for potentiation of the population spike (Taube and Schwartzkroin, 1988). In a study to test a hypothesis put forward by Wilson et al. (1981) that tetanization induces greater LTP of the excitatory pathway than of the feedforward inhibitory pathway in the same area, Abraham et al. (1987) examined the contribution of synaptic inhibition in the E-S potentiation. It was postulated that alteration of the EPSP is not a prerequisite for potentiation of the population spike and that E-S potentiation is largely due to modification of inhibition (Abraham et al., 1987). The possibility of changes in the strength of the inhibitory circuit underlying this overproportional enhancement of the dendritic EPSP-spike relationship led to the suggestion that the term “ L T P should be confined to the changes of the dendritic EPSP (cf. Gustafsson and Wigstrom, 1988). As discussed in Section V,A, tetanization of afferent fibers produces opposite changes of the efficacy of synaptic inhibition: postsynaptically, a reduction of GABA receptor sensitivity; presynaptically, enhanced efficacy of GABAergic transmission through an increase of the number of discharging inhibitory interneurons upon orthodromic stimulation at a given stimulation intensity. Figure 10a demonstrates that a partial block-

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ade of GABA, receptors shortly after the washin of bicuculline results in an increase of all synaptic components including the GABA,-mediated IPSP. Longer exposure to bicuculline (and a more complete block of GABA, receptors) then leads to a progressive reduction of the GABAAmediated IPSP and progressive increasesof EPSP and late IPSP (cf. Fig. 1). These data show that the extent of GABA, receptor blockade determines whether the early IPSP upon orthodromic stimulation of the Schaffer collaterals is enhanced, unchanged, o r decreased. T h e extent of the increase of the EPSP amplitude (and duration) is determined by the extent of GABA, suppression; however, a marked “increase of the E P S P is expressed at any stage (including early stages) of GABA, receptor blockade (cf. Section 111,A). As inferred from the pharmacological blockade of GABA, receptors, the two opposite changes in the inhibitory efficacy after tetanization-suppression of GABA sensitivity and activation of the inhibitory circuit-may not be independent processes: activation of the inhibitory circuit is rather contingent upon reduction of GABA, sensitivity. As for the late, GABA,-mediated IPSP, tetanization results also in a decreased sensitivity of GABA, receptors, thus counteracting the effects of a reduction of GABA, receptor sensitivity following tetanization. When the stimulation intensity of the orthodromic conditioning pulse is adjusted after the tetanus such that the population spike amplitude measured in the CA1 pyramidal cell layer matches pretetanus controls (Fig. 9), “EPSP increases” can still be observed at depolarized potentials including the cell’s resting membrane potential (in the cell depicted about -68 mV) (Fig. 9). This EPSP increase at RMP occurs despite a much smaller EPSP measured at the chloride reversal potential (in the cell depicted at about -78 mV) and at more hyperpolarized membrane potentials. These data suggest that under physiological conditions (cell at resting membrane potential, all excitatory and inhibitory synaptic components preserved) a persistent “enhancement” of synaptic transmission following tetanic stimulation of afferent fibers in the CA1 hippocampal subfield is expressed when the reduction of the efficacy of synaptic inhibition is greater than the reduction of the efficacy of excitatory transmission. An increase of the fast EPSP mediated by AMPA receptors is not a requirement for the manifestation of IXP. It is conceivable that the tetanization-induced increases of intradendritically measured EPSPs are due to the same E-S potentiating mechanisms, i.e., reduction of inhibition (only at a smaller scale at spatially confined areas in distant dendritic regions). Such a notion is supported by the fact that tetanization-induced increases of intradendritically measured EPSPs are small or nonexistent (‘Taubeand Schwartzkroin, 1988).A recent study shows that tetanization of the Schaffer collaterals produced a long-term enhancement of the

239

GABAA RECEPTOR CONTROL POST TET.1

CONTROL

mrac.

Stimul. 1.5 V

Stimul. 2.5 V

Y 40 mo

h

nV

Intrac.

-

MP: 88 m

-98rnV

v

v

LL '

400mo

' zomv

FIG.9. Extracellular field potentials in the CAI pyramidal cell layer (upper row) and intracellular recordings of a CAI pyramidal cell (below). The stimulation strength of the conditioning pulse posttetanum 1 was adjusted to produce a similar population spike amplitude before and after the tetanus. Both, high-frequency tetanic stimuli and the conditioning pulse were administered to fibers in stratum radiatum. Intracellular recordings were performed at the cell's resting membrane potential (-68 mV), at the chloride reversal potential (-78 mV), and at a more hyperpolarized potential at which the GABAAmediated chloride component was reversed and depolarizing (here -98 mV).

compound EPSP in given CA1 pyramidal neurons but no increase in the unitary EPSP elicited by depolarizing current pulses of monosynaptically coupled CA3 neurons (Friedlander et al., 1990). The most probable explanation for these observations (although not taken into consideration by the authors of the study) is that the reduction of the feedforward inhibitory pathway, which involves both GABAA- and GABA,-mediated IPSPs, may have produced the increased dendritic compound EPSP. T h e (unitary) EPSP, however, is not enhanced due to the lack of activation of intercalated inhibitory interneurons in the monosynaptically coupled CA3-CAI neuronal connection. At resting membrane potential, very small increases of subthreshold EPSPs result in cell discharges following tetanization (Figs. 2 and 8). The main physiological mechanisms through which GABA, receptors control the excitability of neuronal populations are large conductance increases in spatially confined subcellular areas (shunting inhibition; cf. Section 111,A). GABAA-mediated potentials elicit only relatively small hyperpolarizing membrane changes at the cell's resting membrane po-

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tential due to the closeness of the chloride reversal potential. The combination of large conductance increases and small membrane potential changes restrict the propagation of GABA,-mediated IPSPs. Tetanization-induced reductions of GABA,-mediated IPSPs generated at far dendritic sites may not be recorded by intracellular recordings in the cell soma, whereas EPSP increases (due to local disinhibition at far dendritic sites, at which a reduction of GABA, sensitivity is effective) can be globally measured (e.g., in the soma) as a potentiated EPSP. It is conceivable that following tetanization, increased EPSPs and “unchanged” fast

A

1,

”.\;

6 Control ............................

Ij 1,

L------

4

Bic(5min) ..............................

2

\

\

40 msec

PT5

FIG.. 10. (A) Orthodroniic potentials of a CAI pyramidal cell during washin of bicuciIlline-methiodide (50 pkf). In comparison to the control response, the early IPSP is initially enhanced ( 5 mill after the start of bicuculline bath application, Bic 5 min), about equal in amplitude (8 miri after bicuculline application, Bic 8 min), and finally completely suppressed (Bic 16 rnin). The EPSP and the late IPSP are potentiated at any point of time following bicuculline application. Cell holding potential -60 mV (by DC current injection). (U) Orthotlromic responses in a CA 1 pyramidal cell in the presence of bicuculline-methiodide (50 pbf) before and at various intervals after tetanization. Large increases in both EPSP and late IPSP occur shortly after application of high-frequency stiniulation to the Schaffer collaterals. Orthodi-oniic responses decrease gradually with time and reach pretetanus control values sonie 20 to 30 min after the tetanus. Recordings were obtained at cell R M P (-68 mV).

GABAA RECEPTOR CONTROL

24 1

IPSPs are recorded in the soma when GABA,-mediated inhibition is effectively suppressed at dendritic sites. The notion that a reduction of fast inhibition represents the major mechanism in the expression of LTP is further supported by the consistent observation that in disinhibited slices long-term enhancement of excitability is not observed (Fig. 10). Tetanic stimulation of afferent fibers after complete blockade of GABA, receptors produces large increases in both EPSP and late IPSP shortly after the tetanus (Fig. 1Ob). However, orthodromic responses decrease gradually with time and reach pretetanus control values about 20 to 30 min after the tetanus (Fig. lob). In summary, GABA, receptors may play a critical role in all phases of LTP: 1. Synchronized bursts of CA3 cells could represent a physiological stimulus capable of producing LTP-like plasticity in CA 1. 2. Impairment of GABA,-mediated inhibition may be involved in the induction of LTP. The resulting block of the feedforward inhibitory pathway may promote membrane depolarization of the CA1 pyramidal required to activate NMDA receptors. 3. Reduction of GABA,-mediated synaptic inhibition due to an impairment of GABA, receptor sensitivity is conceivably the major component of potentiated synaptic transmission following tetanization.

C. INTRACELLULAR REGULATION OF GABA, RECEPTOR FUNCTION A number of recent studies have provided evidence for the concept of neuromodulation via intracellular second messenger-induced phosphorylation of ligand- and voltage-gated ion channels. It is now widely accepted that protein phosphorylation and dephosphorylation of receptor-channel complexes represent the primary mechanisms of controlling the efficacy of a number of voltage- and ligand-gated ion channels in the central nervous system (for review, see Nestler and Greengard, 1984; Schwartz and Greenberg, 1987). Historically, much more attention has been paid to the study of upregulation processes of phosphorylation, which have been considered more highly regulated and more relevant for the control of cellular processes. Yet protein phosphorylation functions as a reversible signaling system and equally effective dephosphorylation of protein kinase target enzymes is required to terminate the responses and maintain responsive phosphorylation systems. The state of phosphorylation of a target protein is determined by competition between a phosphorylating protein kinase and a dephosphorylating phosphatase (for review, see, for exam-

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ple. Krebs and Beavo, 1979; Cohen, 1980, 1989; Klee et al., 1988). Several brain phosphatases have recently been described and dephosphorylation as a signal transduction mechanism has been established in the regulation of many cellular processes in the CNS (Nestler and Greengard, 1984; Klee et al., 1988; Armstrong, 1989). Recent studies indicate that phosphatases are regulated by second messengers and protein modulators: activation is either direct, as in the case of phosphatase-2B, which is activated by calmodulin (Klee et al., 1979; Stewart et al., 1982; Klee and Cohen, 1988), o r indirect, through regulation of other phosphatase inhibitors (e.g., CAMP-dependent DARPP-32), as in the case of phosphatase type 1 (Hemmings et al., 1984; Halpain et al., 1990). 1. PhosphoqdutionlDephosphorylation Regulates GABA, Receptor Function Despite the huge body of information about the extracellular modifiability of the GABA,, receptor function (cf. Olsen and Venter, 1986), little was known about possible intracellular regulatory sites until very recently. The advent of the patch-clamp recording technique, in particular the whole cell-clamp approach, has provided a tool to study intracellular regulation processes in a variety of preparations (cultured neurons, acutely dissociated neurons, brain slices) (cf. Hamill et al., 1981; Edwards et al., 1989). The use of low-resistance electrodes (1-3 M a ) allows a rapid dialysis (within 1 to 2 min) of small diffusible molecules and ions and a proportionally slower exchange of larger molecules and small cell organelles (Marty and Neher, 1983). This property becomes even more prominent in recordings from acutely isolated cells in which the widespread deridritic arborization is largely cut (Kay and Wong, 1986). In acutely isolated CA 1 hippocampal neurons of adult guinea pigs. GABA-mediated chloride currents measured in the whole cell-clamp configuration diminish progressively when the intracellular contents of these neurons are perfused with a “minimal” intracellular solution (130 mM trismethanesulfonate, 10 mM HEPES, 10 mM BAPTA, and 0.1 mM leupeptin, pH 7.3) (Stelzer et al., 1988) (Fig. 11A). The decline of the

FIG.1 1. Rundown of GABA, currents in Mg-ATP-free intracellular perfusate resulting from the loss of GABAA conductance. (A) Whole cell recordings of GABA,, outward currents at I , 3 , and 5. inin after cell penetration (top to bottom). (B) Normalized GABA, currents ( A ) and GABA, conductances (m) decreased with time; cell input resistance ( 0 ) and GABAA reversal potential (+) remained unchanged. (C) rundown o f GABAA currents (recordings at 1, .5, and 10 niin following cell penetration at - 10 mV) and stability of glutamate responses (recordings at 30 sec following respective GABAA currents at -60 mV) (from Stelzer t t al., 1988).

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GABA.,-mediated current is accompanied by a proportionate reduction of the GABA, conductance, whereas the GABA reversal potential and the general condition of the recorded neurons are unaltered (as assessed by parallel measurements of cell input resistance and voltage-gated sodium and glutamate-activated inward currents) (Fig. 11B and C). A similar decline of GABA, currents was observed in hippocampal cultured neurons (Vicini et al., 1986; StelLer et al., 1988) and chick sensory neurons (Gyenes et al., 1988). T h e decline of the GABA,-activated conductance is significantly reduced by adding M g 2 + ions and adenosine triphosphate to the intracellular medium: in the presence of 4 mM intracellular MgCl, and 2 mM ATP, GABA, whole cell currents are maintained to about 95% of the control value after 10 min of cell penetration, whereas in the absence of these "stabilizing" factors, GABA, whole cell currents are suppressed to less than 10% of control values at the same point in time (Stelzer et al., 1988; Chen et al., 1990). T h e intracellular presence of Mg-ATP is necessary but not sufficient to maintain GABA, receptor function in hippocampal pyramidal cells. A loss of [Ca2+jichelators during dialysis (cf. Byerly and Moody, 1984) results in elevations of [Cay+],.In the absence of chemical [Ca2+],buffer in the dialysate, GABA, currents arid conductance diminish over a time course similar t o that in the absence of intracellular Mg-ATP (Fig. 12).

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FIG. 12. Elevated [(;a2+ ]i suppresses GABAA-mediated currents. (A) GABA currents recorded in the presence of high [Gas+], and Mg-ATP. Averaged recordings of four cells; the inset depicts recordings of one cell obtained at 0 . 5 , and 10 inin following cell penetration. (B) Decreasing GABA responses in intracellular solution containing Mg-ATP and high [Ca'+], are reversed upon introduction of low [Ca2+Ii.GABA currents below the graph are recorded at the labeled points of time (from Chen el al., 1990).

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The inclusion of 10 mM BAPTA into the recording pipette is a further requirement to stabilize the GABAA response (Stelzer et al., 1988; Chen et al., 1990). There are at least two possible causes for the time-dependent decrease of the GABA, conductance in the absence of [Ca2+Ii buffer. First, GABA channels could be irreversibly destroyed by proteolysis during whole cell recording. The irreversible process of proteolysis is largely eliminated by providing leupeptin, an inhibitor of [Ca2 Ii-dependent proteases. Most notably, the rundown caused by elevation of [Ca2+Ii (1 mM BAPTA, 1 mM Ca2+, yielding about 1 pA4 free calcium in the dialysate) can be reversed by buffering [Ca2+Iito low levels during the whole cell recording (Fig. 12B). The reversibility of GABAA rundown argues against a mechanism via proteolysis of the GABA, receptorfchannel protein. Rundown caused by intracellular perfusion of a Mg-ATP-free solution can similarly be reversed by reintroducing phosphorylation factors (Mg-ATP) (Chen et al., 1990). T h e requirement for Mg-ATP and low [Ca2+Ii suggests that the GABA, receptor or a closely associated regulatory protein needs to be phosphorylated for the receptor to respond to GABA: in the absence of phosphorylation factors (Mg-ATP) or physiological or chemical [Ca2+Ii buffer, dephosphorylation becomes dominant and causes rundown (Stelzer et al., 1988). T h e results of a series of recent experiments support the notion of a (reversible) phosphorylation-dephosphorylation cycle regulating GABA, receptor function. GABA, current rundown due to prevailing dephosphorylation is demonstrated by intracellular application of the unspecifically dephosphorylating enzyme alkaline phosphatase in the presence of phosphorylating factors (Mg-ATP) (Chen et al., 1990) (Fig. 13). Alkaline phosphatase are nonspecific enzymes that hydrolyze phosphorus-containing compounds (McComb et al., 1979). When intracellular ATP is replaced by its more hydrolysisresistant analog adenosine 5'-0-(3-thiotriphosphate)(ATP-y-S), the alkaline phosphatase-induced decline of GABA currents is significantly retarded (Chen et al., 1990) (Fig. 13).The ATP analog ATP-y-S serves as a substrate for protein kinases and the resulting thiophosphate is far more resistant to hydrolysis by phosphatases than is the corresponding phosphate group (Eckstein, 1985). Similarly,the loss of the GABAA conductance caused by high [Ca2 Ii (cf. Fig. 12A) is significantly slower in the presence of ATP-y-S (cf. Chen et al., 1990). T h e latter observation also militates against the notion that the main intracellular effect of Mg-ATP is to maintain low levels of [Ca2+Ii at the inner surface of the plasma membrane (cf. Byerly and Yazejian, 1986): if the primary effect of Mg-ATP in maintaining GABA, receptor function were to lower [Ca2+Ii,one might expect ATP-y-S to +

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FIG. 13. ATP-7-S antagonizes the effect of alkaline phosphatase of reducing GABAAmediated currents. Alkaline phosphatase was introduced into cells that contained equivalent aniounts of ATP and ATP-7-S, respectively. Control responses are averages of four cells containing Mg-ATP (front Chen el nl., 1990).

enhance rundown of GABA, currents, as it is a poor substitute for ATP in active transport processes. T h e conspicuously slower rate of [Ca2+Iiinduced rundown of GABA, currents in the presence of W-7, a calmodulin inhibitor (cf. Tanaka et al., 1982), indicates that [Ca2+]],-activated GABA, rundown is mediated by calmodulin (Chen et al., 1990) (Fig. 14). Calmodulin-dependent phosphatase, calcineurin (phosphatase-BB), has been isolated from many tissues (Klee et al., 1979, 1988; Stewart et al., 1982; Ymg et al., 1982; Krinks et al., 1988; Farber et al., 1987; for review, see Tallant and Cheung, 1986). Calcineurin is-in contrast to phosphatase- 1, -2A, or -2C-not involved in dephosphorylation of metabolism-controlling proteins, indicating a relative restricted substrate specificity (cf. Ingebritsen and Cohen, 1983; Klee and Cohen, 1988). Immunohistochernical studies demonstrate a highly specific regional and subcellular distribution (cf. Wood et al., 1980; Goto et al., 1986): although calcineurin is present in neurons throughout the brain, a marked regional variation with higher levels in the hippocampal formation, the caudatoputamen, and substantia nigra is observed. Calcineurin is specific for neurons and is not detected in glial cells. Like CaM, calcineurin is enriched in dendrites and the somata. T h e abundance of

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FIG. 14. W-7, a calmodulin inhibitor, retards the high [Ca2+]i-induced rundown of GABA currents. Averages of peak amplitudes of GABA currents are depicted for control (low [Ca*+];),high [Ca*+]i (A),and high [Ca2+]i plus W-7. All intracellular solutions contained Mg-ATP (from Chen et al., 1990).

calcineurin in the CNS, its heterogeneous distribution, and its narrow substrate specificity suggest an important contribution of calcineurin to brain function. Intracellular perfusion with the catalytic subunit of calcineurin results in a rapid rundown of GABA, responses that is retarded by additional inclusion of ATP-y-S or W-7 (Q. X. Chen, A. Stelzer, A. R. Kay, and R. K. S. Wong, unpublished observations). In summary, the results discussed in this section indicate that GABA, receptor function is regulated by a phosphorylation/dephosphorylation cycle. The GABA, receptor or a closely associated regulatory protein can exist in a phosphorylated or a dephosphorylated state. With the provision of Mg-ATP, a protein kinase phosphorylates the molecule and maintains the receptor in a functional form. Phosphorylation competes with a Ca2 /calmodulin/phosphatase-dependentdephosphorylation process that renders the GABA, receptor nonfunctional. A similar phosphorylation-dephosphorylation cycle critical for channel function has been proposed for voltage-activated [Ca2 Ii channels (Doroshenko et al., 1984; Eckert and Chad, 1984; Chad and Eckert, 1986; Hosey et al., 1986; Kameyama et al., 1987; Hescheler et al., 1987; Chad et al., 1987; Byerly and Hagiwara, 1988; Armstrong, 1989). Similar to GABA, receptors, [Ca2+]i/caimodulin-dependent calcineurin renders voltage-activated Ca2 channels nonfunctional (cf. Armstrong, 1989). +

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2. Regulation of GABA, Receptor Function b~ [Ca2+],

Effective intracellular [Cay li buffering systems control the homeostasis of [Ca2+Ii, maintaining low levels of [Ca' +Ii and transmembrane Ca' gradients over four orders of magnitude (for review, see McBurney and ISeering, 1987). Both low levels of [Ca2+J i and high transmembrane <;a'+ gradients are vital for maintaining cell integrity (Nelson and Foltz, 1983; Farber, 198 1). However, changes of [Cay Ii are equally important for a variety of cell functions. The control of magnitude and of temporal and spatial distributions of [Ca2+], represents a complex interplay of various Ca'+ -elevating and buffering systems. It is generally accepted that intracellular calcium [Ca2 Ii represents a major second-messenger system, mediating numerous cellular effects. Elevations of [Ca2+J i , which is at a very low concentration under resting conditions (about 85 nM in CA 1 pyramidal cells (Connor et al., 1988), are mediated by Ca' influx through ligand- or voltage-gated ion channels (McDermott el al., 1986; Regehr et al., 1989) o r through mobilization from intracellular stores by inositol triphosphate (IP3) (cf. Berridge and Irvine, 1984). Immediate targets of [Ca2+Iiare cytosolic and membranebound proteins, including several Ca2 -regulated ion channels (cf. Marty, 1989). Several recent reports support the notion of a crucial role of [Ca2 li in the regulation of GABA, receptor function. In pituitary intermediate lobe cells, spontaneous and GABA-induced chloride currents were restricted to ranges of [Ca2+Iibetween 1 nM and 1 pM [Ca2+Ii(Taleb et uf., 1987). Optimal GABA, responses were obtained near physiological [Ca2 I i reduced open times of chloride channels, and GABA responses were virtually abolished at 1 cul/l [Ca2+li(Taleb et al., 1987). In bullfrog sensory neurons, elevation of [Ca' + I i resulted in a short-term decrease of GABA responses due to a reduced GABA binding affinity to its receptor (Inoue et al., 1986). In a biochemical study, however, it was demonstrated that in synaptic membranes of the rat brain, ["H]GABA binding to the GABA,, recognition site was potentiated by [Ca2+Iiand the increased efficacy of GABA, binding required the presence of calmodulin (Majewska and Chang, 1984). The discrepancy in the findings of both studies concerning the effect of [Ca2+Iion GABA,\ binding may be due to differences between vertebrate and invertebrate preparations or due to different short- and long-term regulation mechanisms: because the [Ca'+ Ii-nietliated effect of decreasing the affinity of the GABA,, receptor is observed only milliseconds after Cay+ influx (Inoue et ul., 1986), [Ca2+Ii effects in this system are more likely directly mediated than indirectly through the activation of a [Ca' Ji-dependentkinase or phosphatase. +

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T h e reversible and fast washout of GABAA responses can be blocked by providing measures promoting intracellular phosphorylation. Elevation of [Ca2+Ii shifts the balance of the GABAA maintaining phosphorylation/dephosphorylation cycle toward dephosphorylation by activating calcium/calmodulin-dependent phosphatases (see above) (Chen et al., 1990). In the hippocampal slice, picrotoxin-sensitive spontaneous IPSPs are suppressed following short trains of action potentials induced by intracellular current injection (5-100 at 20 Hz) (Pitler and Alger, 1990). T h e degree of IPSP suppression was proportional to the degree of stimulation. Inclusion of [Ca2+libuffer (EGTA or BAPTA) in the recording solution prevented the suppression of spontaneous IPSPs, whereas bath application of BAY K 8644, which increases Ca2+ influx, enhanced the effect. These data suggest that physiological fluctuations of [Ca2+Iican modify GABAA-mediated IPSPs. a. Target Proteins of [Ca2+Ii in the Regulation of GABA, Receptor Function. T w o calmodulin-dependent mechanisms have been described in the regulation of GABAA receptor function: [3H]GABA binding to the GABA, recognition site is enhanced by Ca2 (EC,, 0.1- 1 p W ) and the stimulation is dependent on the presence of calmodulin (Majewska and Chang, 1984). A target protein for the calmodulin effect was not identified. Elevation of [Ca2+Iishifts the balance of the GABA,, maintaining the phosphorylation/dephosphorylation cycle toward dephosphorylation by activating calcium/calmodulin-dependent phosphatase-2B (Chen et al., 1990). There is little information so far about a possible role of CaM I1 kinase or calpain in the regulation of GABAA receptor function. A direct phosphorylation of the GABAA receptor by purified CaM kinase I1 could not be measured (Browning et al., 1990). b. Sources of[Ca2 +IiChanges in the Regulation of GABA, Currents. T h e site and source of [Ca2+Iiincreases may represent critical factors in the regulation of GABAA receptor function: increases of [Ca2+Ii through voltage-gated calcium channels in bullfrog ganglion cells reduce GABAbinding affinity (Inoue et al., 1986). The amount of suppression of the GABA receptor sensitivity is directly correlated to the amount of Ca2+ influx (maximal for Ca2+ influxes exceeding 80 pC) and is most significant between 10 and 600 msec following voltage-activated Ca spikes (Inoue et al., 1986). The suppression of picrotoxin-sensitive spontaneous IPSPs due to increases in [Ca2+Iifollowing short trains of action potentials during intracellular current injection is also mediated by voltagegated [Ca2+Ii channels (Pitler and Alger, 1990). The tetanizationinduced block of both phases of IPSPs, the early, GABAA-mediated and late, GABA,-mediated IPSPs, in hippocampal slices is prevented by the NMDA receptor antagonist D-APV, indicating that suppression of GABA-mediated IPSPs is contingent upon NMDA receptor activation +

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(Stelzer et al., 198i). There is no direct evidence yet that the hydrolysis of phosphatidyliriositoI-4,5-biphosphateand formation of IP3, resulting in mobilization of [Ca2 J,, regulates GABA, receptor function. I n summary, the question as to whether GABA, receptor function is affected by changes in [Ca'+ 1, may largely be determined by the site and nature of [Can+],gradients produced by various sources of [Ca2+],fluctuations. T h e concept of [Ca2 1, concentration gradients or spatial distribution of [Ca' 1, as a mechanism for signal transduction specification has been implicated in all systems that promote increases of [Ca2+], (cf. Berridge and Taylor, 1988; Harootunian et al., 1988). Even subtle changes of [Cay 1, may exert profound effects in the immediate vicinity of Ca' +-generating sources (ion channels or organelles). Experimental data of the possible subcellular colocalization or of the vicinity of GABA, receptors and Ca2 +-elevating systems (calcium channels, IP3mediated mobilization of [Ca2+I,) are lacking. Suppression of GABA,,mediated IPSPs by short trains of action potentials (Pitler and Alger, 1990) suggests that relatively small [Ca2 J, elevations under physiological conditions may be effective. [Ca2 1, gradients generated by tetanization (in regard to both amount and spatial propagation; cf. Regehr et al., 1989; Regehr and 'IBnk, 1990) could lead to profound suppression of GABA,, receptor function in CA 1 pyramidal cells. +

+

+

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3. Proteiri Kinuse5

Specific forms of the three main protein kinases, Ca'+ /phospholipid-dependent kinase [protein kinase C (PKC)], CAMP-dependent kinase [protein kinase A (PKA)], and Ca' + /calmoddin-dependent kinase (CaM I1 kinase) exist in the central nervous system in high concentrations: PKC arid PKA constitute 0.01-0.1% of total protein (Hart1 and Rokoski, 1982; Kikkawa et ul., 1982) and CaM I1 kinase, u p to 1.4% in selected brain areas (Erondu and Kennedy, 1985).T h e protein kinases are phosphoryltransferases, catalyzing the transfer of the terminal phosphoryl group of ATP to substrate proteins at selected serine and threonine residues (cf. Nestler and Greengard, 1984; Schwartz and Greenberg, 1987). T h e amino acid sequence in the vicinity of the acceptor hydrox): group determines the (broad) substrate specificity (cf. Nestler and Greengard, 1984). Structural and biochemical studies have proposed several phosphorylation sites at various subunits of the GABA, receptor: a CAMPdependent phosphorylation site at the p, subunit (Schofield et al., 1987; Browning et al., 1990; Pritchett et al., 1989),a PKC phosphorylation site at the p subunit (Browning et al., 1990),a tyrosine kinase site (Pritchett t,t

GABAA RECEPTOR CONTROL

25 1

al., 1989) at the y2 subunit, and an unspecified phosphorylation site at the a subunit (Sweetnam et al., 1988). a. A n Endogenous Protein Kinase Phosphorylates the ci Subunit of the GABA, Receptor. Sweetnam et al. (1988) provided the first evidence that the (Y subunit of the GABAA receptor is phosphorylated in partially purified preparations of the GABA,/benzodiazepine receptor from rat brain. T h e a subunit phosphorylation of the GABAA receptor was demonstrated by the comigration of the phosphorylated protein on twodimensional gels with photolabeled, immunolabeled, and purified a subunit and by identical peptide maps generated by the phosphorylated protein and iodinated, purified ci subunit. Evidence was provided that an endogenous protein kinase in the crude brain membrane preparation catalyzed the observed phosphorylation of the GABA, receptor ci subunit. However, the protein kinase could not be activated by CAMP, cGMP, calcium alone, calcium plus calmodulin, or calcium plus phosphatidylserine, nor inhibited by specific inhibitors of CAMP-, cGMP-, calcium/calmodulin-, or calcium/ phosphatidylserine-dependentkinases. T h e authors concluded that the ci subunit phosphorylating protein kinase might be second-messenger independent. b. Protein Kinase A and C Phosphorylate f3 Subunits of the GABAIBenzodiazepine Receptor. Kirkness et al. ( 1989) demonstrated protein kinase A-dependent phosphorylation of the p subunit of the GABAA receptor: in a preparation of GABAIbenzodiazepine receptors from pig cerebral cortex, which consists of three major bands of polypeptides (5 1,55, and 57 kDa in a ratio 2: 1:l), treatment with CAMP-dependent kinase resulted in major incorporation of 32Pinto the 55-kDA band, a muscimol-binding polypeptide (p subunit). Browning et al. (1990) also reported the phosphorylation of the @ subunit by CAMP-dependent protein kinase: protein kinase A, purified from bovine heart, phosphorylated a polypeptide band of the purified GABAA receptor (M, 58,000), termed p5*. Purified protein kinase C phosphorylated a polypeptide band (p56) with an apparent M , of 56,000. Comigration of the phosphorylated bands with muscimol-binding polypeptides was confirmed. Polypeptides labeled with the p subunit-specific antibody incorporated significantly less 32P following application of PKA or PKC, indicating that the p subunits of the GABA, receptors are the main target proteins of phosphorylation by protein kinase C and A. In this study, CaM kinase I1 did not produce a significant phosphorylation of the GABAA receptor. However, CaM I1 kinase enzyme is the main target of CaM II-mediated phosphorylation. T h e autophosphorylation of CaM I1 may have masked net phosphorylation of the purified GABA, receptor (Browning et al., 1990). In summary, the recent biochemical studies by Sweetnam et al.,

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Kirkness et al., and Browning et al. demonstrate conclusively that the major subunits of GABA,\ receptors fie., OL and P subunits) are targets of phosphorylation processes. Protein kinase A and C phosphorylate p subunits, the kinase has not yet been identified. c. GABA,4 Receptor Function: CAMP und Protein Kinase A . Most of the actions of the second-messenger CAMP (for review, cf. Robinson et ul., 1971) in mammalian tissues take place through the interaction of CAMP with the regulatory subunit of cyclic CAMP-dependent protein kinase (EC 2.7.1.37; ATP protein phosphotransferase, protein kinase A) (for review, see Krebs, 1975). Intensive investigations revealed the detailed pathways and proteins involved in the generation of cyclic AMP and subsequent activation of CAMP-dependent kinase. Activation of protein kinase A in response to hormones, neurotransmitters, and other stimulating agents results in protein phosphorylation of many target proteins that have been implicated in a variety of cellular processes, including ion channel modulation, transmitter synthesis, regulation of growth, and gene transcription (for review, see Krebs and Beavo, 1979; Nestler and Greengard, 1984). The phosphorylation catalyzed by PKA can be reversed by dephosphorylation processes catalyzed by specific phosphatases (cf. Klee and Cohen, 1988; cf. discussion). In 1971, Walsh et al. described a heat-stable protein in rabbit skeletal muscle; this protein inhibited protein kinase A in a competitive and specific manner by binding to the catalJ4c subunit of PKA (& = 1 nM; Demaille et al., 1977). The protein kinase A-inhibiting enzyme can be found in many tissues, including brain (Ashby and Walsh, 1973; Beavo and Mumby, 1982). Estirriates suggest that about 20% of the total protein kinase activity in skeletal muscle can be inhibited by the protein kinase inhibitor present in ili710 (Wdlsh and .4shby, 1973; Beavo and Mumby, 1982). Several recent studies provide compelling evidence for CAMP-dependent phosphorylation of the GABA, receptor. First, it was shown that the pL' subunit Lf the bovine GABA, receptor contains a unique CAMPdependent serine phosphorylation consensus sequence (Schofield et al., 1987; Pritchett Pt ul., 1989). Second, actual phosphorylation of the p subunit of the purified GABA, receptor by CAMP-dependent kinase has been demonstrated in receptor preparations from pig cerebral cortex (Kirkness et a/., 1989) and from rat neocortex (Browning ot nl., 1990). Reports about possible functional consequences of CAMP-dependent phosphorylation of the GABA,4 receptor are controversial: CAMP arid the adenylate cyclase stimulator forskolin decreased GABA-gated chloride fluxes in rat brain synaptoneurosomes (Heuschneider and Schwartz, 1989). A similar reduction of GABA-mediated currents was

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observed in chick cortical neurons following administration of forskolin and the cAMP analog 8-bromo-CAMP(Tehrani et al., 1989). In addition, a fast component of desensitization appeared following the application of PKA stimulators (Tehrani et al., 1989). In mouse spinal neurons, inclusion of the purified catalytic subunit of PKA (50 pMlml in the pipette solution reduced the peak conductance of whole cell currents evoked by GABA (Porter et al., 1989). Two other features of GABAA receptors (desensitization and binding) have been reported to be modified by the cAMP/PKA system: In chick cortical neurons, both 8-bromo-CAMPand forskolin reveal a fast desensitization component, an effect accompanied by a reduction of the peak current amplitude (Tehrani et al., 1989). Biochemical studies show that a heat-stable protein inhibiting GABA binding (GABA modulin) loses its inhibitory efficacy through CAMP-dependent phosphorylation (Wise et al., 1983): CAMP-dependent phosphorylation of GABA modulin would enhance the efficacy of GABA binding. Several effects by agents commonly used to stimulate the cAMP second-messenger system are not related to PKA-mediated phosphorylation: cAMP and cAMP analogs (1-2 mM) decrease GABA, responses through mechanisms not related to stimulation of PKA (Harrison and Lambert, 1989). In addition, a direct interaction of forskolin with the chloride channel was proposed, because 1,9-dideoxyforskoIin, which does not activate adenylate cyclase, also inhibited the muscimol-stimulated chloride uptake in synaptoneurosomes (Heuschneider and Schwartz, 1989). Forskolin and cyclic AMP analogs (8-bromo-adenosine, 8-bromo-adenosine CAMP,dibutyryl CAMP,and CAMP)inhibited GABAinduced 36C1- influx in mice spinal cord cultured neurons without the involvement of protein kinase A (Mehta and Ticku, 1990). Muscimolstimulated 36Cl- uptake in mouse brain microsacs was inhibited by cAMP analogs (CAMP, 8-bromo-cAMP, CPTcAMP) in a millimolar range (Leidenheimer et al., 1990). Pretreatment with the PKA inhibitor H-8 did not affect CAMP-mediated impairment of muscimol-stimulated 36Cluptake, indicating that the effect is phosphorylation independent. The cAMP analogs (CAMP,8-bromo-cAMP, and CPTcAMP) at low millimolar concentrations and H-7 inhibited binding of the GABA, receptor ligand t3H]SR 9553 1, whereas phosphodiesterase inhibitor IBMX and adenylate cyclase activator forskolin inhibited binding of the C1- channel ligand [3%]TBPS (Leidenheimer et al., 1990). In dissociated CA 1 hippocampal pyramidal cells, 8-bromo-CAMP produces concentration-dependent effects on GABAA currents: bathapplied 8-bromo-CAMP at concentrations higher than 50 pM results

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in a monophasic reduction of GABA, currents. T h e suppression of GABA, currents is mimicked by extracellular cAMP at these concentrations. Low niicroniolar levels of 8-bromo-CAMP (1-10 p M ) result in a biphasic alteration of GABA, peak-current amplitudes: a transient increase of GABA, currents is followed by a marked decrease. Extracellular CAMP in this concentration range produces a monophasic reduction i n some cells and no changes of GABA,\ responses in others. At nanomolar concentrations of 8-brorno-CAMP( 10- 100 nM), however, monophasic potentiating GABA, changes were observed in about half of the neurons recorded (Fig. 15). Extracellularly applied cAMP at such low levels does not affect GABA, responses. 'These data suggest that the specific effect of S-bromo-CAMP (i.e., not mimicked by extracellular CAMP)is a potentiation of GABA, currents. 'The notion that the cAMPIPKA system up-regulates GABA, receptor function is supported by the observation that intracellular application of Walsh PKA inhibitor (Peninsula 5-24, 20 nM) caused the GABA, responses in isolated hippocampal cells to run down in the presence of stabilizing factors Mg-ATP and BAPTA (Fig. 15). The time course of rundown in the presence of PKA inhibitor is similar to that observed in the absence of phosphorylation factors or in the presence of high [Caz +I,. (1. GABA, Rereptor Fmctzon: Prolein Kinase C. Since protein kinase C (PKC) was discovered in 1977, a number of studies have established its prominent role in cellular signal transduction (for review, see Nishizuka, 1986, 1988, 1989; Huang, 1989). PKC is physiologically activated by the second messenger diacylglycerol (DAG), a breakdown product of the phospholipase C-mediated hydrolysis of the membrane phospholipid phosphatidylinositol-4,5-biphosphate.Small amounts of' DAG increase the affinity of PKC for [Ca'+], present and activate the emyme without changes of [Ca"+], (Wolf el al., 1985). PKC was shown by quantitative receptor autoradiography to bind [3H]phorbol-12,1%dibutyrate, a tumor-promoting stimulator of PKC, with high affinity and thus to be localized as enzyme in the brain (Worley et id., 1986). It is enriched in certain brain regions, with the highest concentration in the CA 1 hippoFIG.1.5. (Xtp) Extracellular application of 10 n b f 8-bromo-CAMPproduces an increase of <;ABA-mcdiated chloride currents in acutely dissociated C A I pyramidal cells in the presence ctf intracellular Mg-ATP (holding potential, - 10 mV). (Bottom) Intracellular inclusion of protein kinase A inhibitor (Peninsula 5-24, 20 nhf) results in rundown of C A R A A whole cell currents (m, average of eight cells). Averaged control GABAA responses ( + , ) I = I I ) under stabilizing conditions are depicted for comparison.

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campal subfield (Worley et al., 1986). Within the CA1 hippocampus, highest concentrations of PKC are found in the cell body layer of stratum pyramidale (Roth et ai., 1989). Among the PKC subspecies, the y and the two p forms are enriched in stratum pyramidale (Brandt et al., 1988). In the CIVS, PKC has been implicated in the regulation of transmitter release, ion channels, plasticity, and cell development. The function of a number of voltage- and ligand-gated channels can be modified by PKC-mediated phosphorylation (Baraban et al., 1985; DeRiemer et a / . , 1985; Kane and Dunlap, 1986; Holz et al., 1986; Doerner et al., 1988; Lacerda et al., 1988; Sigel and Baur, 1988). Phosphorylation of the GABA,4 receptor by PKC has been demonstrated in biochemical studies: purified PKC from bovine heart phosphorylates mainly the p subunit of the purified GABA, receptor from rat neocortex (Browning et al., 1990). Several potent tools for a functional investigation of PKC effects exist: some synthetic diacylglycerols, such as 1-oleolyl-2-acetylglycerol(OAG), and tumor-promoting phorbol ester are able to penetrate intact cell membranes and activate PKC directly (Kaibuchi et al., 1982; Castagna et al., 1982). In acutely isolated CAI hippocampal neurons o f adult guinea pigs, bath application of nanomolar concentrations of the synthetic diacylglycerol analog 1 -oleolyl-2-acetylglycerol resulted in a steady, progressive reduction of GABA, currents after a few minutes of experimental intervention (Stelzer and M h g , 1988; Stelzer et af., 1988) (Fig. 16). Phorbol ester phorbol- 12,13-dibutyrate (PDBu) mimics the action of OAG on GABA currents in a similar concentration range. Application of the biologically inactive (Y analog of PDBu does not promote such an effect. At lo\\ PDBu or OAG concentrations (up to 250 nM), the decline of the GABA peak-current amplitude progressed to a finite value of about 50%, e\en in the presence of the PKC-stimulating agent. At higher concentration ranges (400 and 500 IM,respectively), application of both types of PKC activators generally leads to a continuous reduction of the GABA response in the presence of the PKC-activating drug, resulting in a complete abolition after sufficiently long administration (Fig. 16). The decline could, however, be brought to a halt at any level by switching back to control saline. These data suggest that PKC activation can virtually abolish the responsiveness of the GABA,\ receptor. A reduction of GABA sensitivity after phorbol ester application was also found in Xenopiu oocytes, in which receptors and ion channels are expressed following the injection of chick brain mRKA (Sigel and Baur, 1988). These data suggest that activation of PKC results in an impairment of GABA,q receptor efficacy. Modulators of PKC at higher concentrations have side effects that

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0

10

20

30

lminl FIG. 16. Application of the synthetic protein kinase C activator OAG (400 “M) results in a suppression of GABAA peak-current amplitudes. Recordings were obtained under stabilizing standard conditions (at - 10 mV, in the presence of intracellular Mg-ATP and BAPTA).

are unrelated to PKC activation (cf. Hockberger et al., 1989, for calcium currents): in acutely dissociated pyramidal cells, PDBu and OAG at concentrations higher than 0.5 and 1 ~LM, respectively, decreased the cell input resistance and shifted the current baseline in an inward direction. Both actions were regularly accompanied by a rapid rundown of GABA, chloride currents. The rapid suppression of GABA, currents at these concentrations is most likely not due to PKC activation because similar effects (changes of passive cell membrane properties and rapid decline of GABA, currents) were observed using the ct analog of PDBu at these higher concentrations.

D. DISCUSSION 1. Intrucellulur R e p l a t i o n of GABA, Receptors

The modulation of GABA, receptor sensitivity via intracellular second messengers provides mechanisms for lasting modifications of the state of excitability of neuronal populations. Data discussed above indicate that GABA, receptor function may be modified by several intracellular agents contributing to a complex interplay of extra- and

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intracellular regulation processes. Based on studies in the hippocampal slice and acutely dissociated neurons (data discussed above), the following scheme for second-messenger regulation of GABA, receptor function in CAI pyramidal cells is proposed (see Fig. 17 for illustration): the amount of GABA, conductance is determined by a competition of phosphorylation, which maintains GABA, conductance, and a phosphatasedependent dephosphorylation, which reduces GABA, conductance (cf. Figs. 11-14). GABA, receptors are phosphorylated at at least two different sites by protein kinase A and protein kinase C, respectively (Sweetnam et ai., 1988; Kirkness et al., 1989; Browning et al., 1990).Stimulation of protein kinase C leads to a suppression of GABA, sensitivity (Fig. 16). T h e two second-messenger systems, [Ca2 1, and PKC, generally believed to procluce long-term changes of excitability in cortical populations, may suppress GABA., receptor function via phosphorylation (PKC) and dephosphorviation ([Ca' ll) at two distinct phosphorylation sites. Protein kinase A maintains, rather than suppresses, GABAA receptor function (Fig. 15). +

+

i t

Calcium channels

PKC

lntracellular

0

c

v

IC.'? 1

GABAA

PI-+

I

Calcineurin]aCaM

I

FIG. 17. Scheme of intracellular regulation of GABAA receptor function (see text).

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GABA, RECEPTOR CONTROL

Inhibition of CAMP-dependent kinase by intracellular application of the Walsh inhibitor causes a loss of GABA, conductance within a few minutes following the establishment of whole cell recordings in CA1 pyramidal neurons (Fig. 15). Considering the effects of PKA inhibition and regulatory features of the CAMP-dependent protein kinase system outlined above, it is conceivable that the primary effect of CAMP-dependent phosphorylation on GABA, receptors is the maintenance of receptor function. A cyclic CAMPmechanism of maintaining GABAA receptor function would conceivably set the stage for a possible antagonism between protein kinase A-dependent phosphorylation and Ca2 /CaM/ calcineurin-dependent dephosphorylation of GABAA receptors. Similar phosphorylation-dephosphorylation cycles have been reported for several other phosphoproteins in the CNS: the antagonism of CAMPdependent phosphorylation and calcineurin-dependent dephosphorylation of dopamine- and CAMP-regulated phosphoprotein (DARPP) (Hemmings et al., 1984; Halpain et al., 1990), the R subunit of protein kinase A (Blumenthal et al., 1986), and microtubuline-associated protein 2 (MAP2) (Halpain and Greengard, 1990) has led to the proposal of a general principle of such phosphorylation-dephosphorylation cycle. Opposition of [Ca*+Ii-and CAMP-dependent signals extends to a multifunctional scheme of interaction beyond the phosphorylation-dephosphorylation antagonism at the phosphoprotein phosphorylation site (cf. Klee and Cohen, 1988; Cohen, 1989). +

2. Tetanization-Induced Progression of Excitability Repeated tetanization of afferent fibers results in a progressive enhancement of excitability in the dentate gyrus in vivo (Bliss and Loemo, 1973) and in CA3 (Stasheff et al., 1985; Miles and Wong, 1987b) and in CAI in vitro (Slater et al., 1985;Stelzer et al., 1987). The progressive increase of excitability following repeated tetanization of afferent fibers represents the gradual transition from physiological resting activity to LTP, which is widely believed to represent an increased state of excitability within “physiological” limits (as cellular substrates of learning processes) to the final stages of highly synchronized epileptiform activity in the form of interictal-like events (paroxysmal depolarization shifts) and electroencephalographic seizures. LTP has been proposed as a cellular substrate for both memory (Swanson et al., 1982) and kindlinginduced epileptogenesis (Racine, 1978; McNamara et al., 1980; Slater et al., 1985). LTP, particularly LTP elicited in vivo, is not an all-or-none event, but rather comprises a wide range of states of excitability (cf. Bliss and Loemo, 1973) considered “within physiological limits.” a. Progressive Excitability Due to Disinhibition. Tetanization-induced

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progression of excitability is accompanied by characteristic alterations of synaptic transmission and emergent properties of the network. T h e changes in the synaptic properties of GABAergic inhibition leading to functional disinhibition (loss of the postsynaptic GABA sensitivity) may account for most of the excitability changes following tetanization (cf. Sections V,A and V,B). In the hippocampal CA3 subfield, emergent circuit properties evoked by tetanization are strikingly similar to those observed following the pharmacological blockade of GABA, receptor function (Miles and Wong, 1987b). Tetanization-induced alterations in synaptic transmission and emergent network properties are due to impairment of inhibitory efficacy, most probably postsynaptic GABA sensitivity (Miles and Wong, 1987b; Stelzer et d.,1987). Enhanced interneuron excitability following tetanization may mask the concomitant reduction of the postsynaptic sensitivity under certain conditions. T h e tetanization-induced decrease in the postsynaptic GABA sensitivity can be assessed by the response to exogenous GABA application, which represents a pure postsynaptic component: GABA sensitivity is reduced as a result of the tetanus (cf. Figs. 5 and 8). Repeated application of tetani to afferents leads to a progressive impairment of GABA receptor sensitivity superseding increases of interneuron activity. At these advanced stages of excitability, all parameters of synaptic inhibition, including orthodromically evoked I PSPS, are generally reduced (cf. Misgeld and Klee, 1984; Abraham et al., 1987; Stelzer et al., 1987). Besides progression of tetanization-induced alterations of synaptic transmission, t w o other general properties of tetanization-induced rnodification of synaptic transmission are prominent: first, the long duration of effects (always until the end of the experiment, up to 4 hr of intracellular recording; cf. Stelzer et al., 1987); second, all tetanization-induced modifications of synaptic transmission and receptor sensitivity resulting in an overall increase of excitability of the CA1 neuronal population are prevented when the tetanus is applied in the presence of the NMDA receptor antagonist D-APV (cf. Figs. 3 and 5). Based on a variety of experimental data (see above) it is postulated that Ca' influxes through NMDA receptor channels represent the causal mechanisms of persistent hyperexcitability in cortical populations, through a reduction of GABA, receptor sensitivity. Possible mechanisms are discussed in the following sections. 6. NMDA-GABA, Receptor Interaction. NMDA receptors have been implicated in long-lasting functional (cf. Collingridge and Bliss, 1987; Cotman and Iversen, 1987) and morphological plasticity (Cotman and Iversen, 1987; Pearce et al., 1987; Balazs et al., 1988; Mattson et al., 1988). NMDA receptors are activated by binding of glutamate, the main +

GABAA RECEPTOR CONTROL

26 1

excitatory transmitter in the mammalian cortex (cf. Watkins and Evans, 1981). Glutamate activates cationic conductances via at least two receptor subtypes: AMPA receptors mediate fast EPSPs and NMDA receptors mediate slow EPSPs (cf. Watkins et al., 1990). c. [Ca2 +Ii and NMDA. The requirement of elevations of [Ca2+Iiand the prevention of functional or morphological changes by NMDA receptor antagonists (e.g., the expression of LTP in the CA1 hippocampal subfield) strongly suggest a causal link between Ca2+ influxes through NMDA receptor-coupled ionophores and the expression of synaptic plasticity (cf. Malenka et al., 1989; Kennedy, 1988). The high Ca2+ permeability for NMDA receptor channels (McDermott et al., 1986; Mayer and Westbrook, 1987; Huettner, 1988; Vyklicky et al., 1988; Regehr and Tank, 1990) in combination with Ca2 ions triggering consecutive second-messenger actions may explain how multiple plasticity effects could be produced by a single class of postsynaptic receptors. A number of intracellular [Ca*+Iitarget proteins have been demonstrated to be activated by NMDA: NMDA stimulates calcium-dependent protease calpain in hippocampal slices (Seubert et al., 1988) and in intact rats (Siman and Noszek, 1988). NMDA-mediated stimulation of calcium/CaM-dependent protein kinase has been proposed as a biochemical mechanism for the maintenance of LTP (for review, see Kennedy, 1989). Calcineurin-mediated dephosphorylation of DARPP-32 and MAP2 is achieved through NMDA receptor activation (Halpain et al., 1990; Halpain and Greengard, 1990). However, direct proof for an actual link between NMDA-mediated [Ca2+Ii influx and consecutive biochemical o r physiological effects is sparse. The investigation of NMDA receptor-mediated [Ca2+Ii effects on GABAA under whole cell-clamp conditions in dissociated neurons poses several experimental problems. First, [Ca2+Iibuffering needs to be effective enough to produce stable GABA control responses, but also permissive enough to allow Ca2 fluxes to produce significant alterations of [Ca2+Ii gradients. As inferred from invertebrate neurons, washout of physiological [Ca2 Ii buffering systems and sequestration systems is difficult to compensate by adding chemical buffers to the dialysate (Byerly and Moody, 1984). Second, recent studies show that the NMDA receptor is sensitive to the action of proteolytic enzyme applied extracellularly (Allen et al., 1988; Akaike et al., 1988). A milder enzymatic dissociation procedure, however, can preserve NMDA responses with typical regulatory features (sensitive to Mg2+, APV, and glycine) (Kay and Connor, 1990). Third, excitatory amino acids play a multifunctional role in GABAA receptor regulation. A [Ca2+Ii-independent potentiation of GABA, whole cell currents by excitatory amino acids (including NMDA) requires lower +

+

+

262

ARhlIN STELZEK

levels of agonists (Stelzer and Wong, 1989) and may mask the effect of NMDA receptor-mediated suppression of GABA, responses. APV prevents the reduction of GABAergic IPSPs and GABA sensitivity, including all GABA, components in CAI pyramidal cells, following tetanization, but picrotoxin-sensitive spontaneous IPSPs are also suppressed by voltage-gated Cay+ currents in CA1 pyramidal cells (Pitler and Alger, 1990). In addition, the expression of LTP, which is contingent upon NMDA receptor activation in some brain areas, including the CA1 hippocampal subfield, occurs in others in a NMDA receptor-independent fashion. The specificity and the persistence of effects triggered by NMDA receptors cannot be explained by the high [Ca2+Ii permeability of NMDA receptors alone. Additional properties of NMDA receptors may contribute to specificity and maintenance of persistent increase of synaptic transmission or other plasticity effects triggered by NMDA receptors. In summary, the bulk of expressions of synaptic plasticity following tetanization of afferent pathways may be carried by the modification of GABA, receptor function, the impairment of which can explain most of the emergent changes of synaptic potentials and network properties, including the potentiation of excitatory synaptic efficacy in LTP. The two second-messenger systems, [Ca' +Ii and protein kinase C activation, generally believed to induce and maintain enhancement of excitability in cortical populations may suppress GABAA receptor function via phosphorylation (PKC) (Fig. 16) and dephosphorylation ([Ca2+Ii)(Figs. 1214) at t w o distinct phosphorylation sites (Fig. 17). The repeated application of tetani may produce two effects on GABA, receptor sensitivity. First, there is a progressive suppression of CABA, receptor function (through increased [Ca' + I i concentrations or other mechanisms discussed above) at sites that had been affected to a lesser extent by a single tetanus. Second, there is a spatial propagation of GABA,-suppressing [Ca' + I i gradients. Large [Cay+ I i gradients may develop in a spatially confined area (e.g., at dendritic shafts): relatively small amounts of Ca2 influx through NMDA receptors could result in large concentration changes in such small compartmentalized areas. Repeated tetanization results in spatial propagation of [Cay ]; gradients and could affect CABA,\ receptor sites that had not been af€ected by a single tetanus. Microfluorometric [Ca2+ I i measurements demonstrate a characteristic subcellular distribution pattern of [Ca' + I i increases following high-frequency stimulation. 'Trains of tetani evoke transient elevations of [Ca' 1; to about 1 phf in apical and basal dendrites of CA 1 pyramidal cells in the guinea pig hippocampal slice preparation (Regehr et al., 1989). After a single tetanus producing LTP, specific NMDA recep+

+

+

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GABAA RECEPTOR CONTROL

tor-mediated increases of [Ca2+Iireach levels of 500 nkf [Ca2+Iiin spatially confined dendritic areas near activated afferent fibers (Regehr and Tank, 1990). T h e tetanization-induced opposite pre- and postsynaptic modifications on synaptic inhibition may lead to a spatial shift of inhibitory synaptic efficacy: after weak tetanic stimulation, GABAA sensitivity will be impaired at sites close to the activated afferents at which [Ca2+Ii gradients can transiently supercede resting [Ca2+Ii levels by magnitudes of several hundreds (Regehr and Tank, 1990). Synaptic efficacy will, however, be more effective at other, more distant sites on the cell surface, which receive inputs from previously silent interneurons recruited by the stronger activation of the feedback inhibitory pathway after tetanic stimulation. With repeated tetanization and intracellular propagation of [Ca*+Ii gradients, more and more GABA, receptors at more distant sites (with respect to the site of actual Ca2 entry) become unfunctional. T h e direction of propagation of reduced GABA, sensitivity is likely to be from distant dendritic to more proximal (somatic) sites. At this stage, a “decrease” in orthodromically evoked IPSPs following tetanization becomes manifest, because the impairment of GABA, receptor sensitivity is comparatively larger (both spatially and in regard to the absolute amount of lost GABA, sensitivity) than compensating presynaptically mediated increases of the inhibitory strength (cf. Section V,B,3). I n addition, the loss of GABA, sensitivity at subcellular locations more proximal to the somatic site will reveal changes of IPSPs in somatic intracellular recordings due to a shorter electrotonic distance. I n summary, the synopsis of experimental observations discussed in this section may allow the conclusion that the NMDA-dependent [Ca2+Ii gradients generated by tetanization of the Schaffer collateral afferents reflect both the amount and subcellular distribution of impaired GABAA receptor function and the general state of excitability of the CAl neuronal population. +

VI. GABAA Receptor Function: Synchronization

T h e generation and propagation of synchronized population activity constitute complex phenomena involving intrinsic features of single cells (such as burst discharges), synaptic processes, circuit properties, and mechanisms such as the generation of a rhythm of synchronized population activity that cannot be sufficiently explained by either intrinsic or synaptic properties alone, or in combination (cf. Traub et al., 1989a). New techniques and preparations, in particular the brain slice (cf.

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Dingledine, 1984) and computer models (cf. Traub et al., 1987), have provided powerful tools to study synchronization properties. T h e results of recent investigations demonstrate the significance of synchronized activity under physiological and pathological conditions. GABA, receptors play a paramount role in the genesis and expression of synchronization at virtually every level, from modifying intrinsic currents producing burst discharges to the activation of previously silent connections and the entrainment of the rhythm of population oscillations.

SYNCHRONIZED ACTIVITY A. PHYSIOLOGICAL The synchronous firing of a large group of neurons and the development of a rhythm of synchronous activity are defining features of epilepti form activity in the central nervous system. Fully synchronized burst activities with inhibition blocked (e.g., in the presence of GABA, antagonists) (Fig. 18) or with inhibition increased and fully synchronized (e.g., in the presence of 4-AP) represent extreme (and pathological) states of the collective behavior of neuronal populations. However, partially rhythmic synchronized activity can be observed in many systems in the nervous systems as an expression of regular rather than epileptogenic brain activity. Rhythmical waves in the electroencephalogram and a variety of complex behaviors are produced by rhythmic oscillations of a large assembly of neurons (Schwartzkroin and Knowles, 1984; Schwartzkroin and Haglund, 1986; Schneiderman, 1986; Miles and Wong, 1978a; Traub et a!., 1989a,b). In the normal resting slice, CA3 neurons generate periodic spontaneous discharges (Wong and Prince, 1981), most of which occur at different frequencies in different cells and out of phase with one another-as shown by the lack of measurable field potential in slice recordings and by the lack of synchronization in pairs of' cells recorded simultaneously with intracellular electrodes. Due to the powerful inhibitory control of the recurrent pathways in CA3, burst propagation between CA3 pyramidal cells is effectively restricted. Inhibitory activity is locally driven by bursts of excitatory cells: the synaptic connectivity and transmission kinetics of excitatory synapses activating inhibitory interneurons provide an effective feedback system for the spatial and temporal limitation of excitatory bursts (cf. Miles and Wong, 1987a; Traub et al., 1989a,b). Simulation studies, however, show that the inhibitory control over the recurrent excitatory pathway under physiological conditions is not complete and burst firing of pyramidal cells can propagate to a certain extent within a confined area of the network (Traub et al., 1989b). Although most of the spontaneous burst discharges

265

GABAA RECEPTOR CONTROL

PTXt=Omin 1

1

%2

B

16 min

1 m

8

C

m

m m

m

D

12mV

I 2 mV m m

m m

m

!

0.5 s E

16min

23 min

27 min

100 ms FIG. 18. Rhythmic synchronous synaptic events precede full synchrony as GABAAmediated synaptic inhibition is suppressed. (A-D) simultaneous intracellular recordings of' CA3 pyramidal cells at various points of time following the application of picrotoxin (4 pM). Filled squares mark synchronous synaptic events which progressively grow in amplitude and frequency. Large spontaneous synaptic events at 27 and 29 min are followed by a hyperpolarization. Rhythmic activity is established at an interval of about 2 sec. (E) Synchronous synaptic events become larger and more complex with time following picrotoxin exposure (from Miles and Wong, 1987a; with permission).

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are out of phase with each other, some spontaneous (partially) synchronized activity, both depolarizing and hyperpolarizing, can be measured in the hippocampus by intracellular recordings from pairs of cells or by recording apical dendritic field potentials (Schneiderman, 1986; Schwartzkroin and Haglund, 1986; Miles and Wong, 1987a; cf. Traub et ul., 1989a.b). Although the basic circuitry of the hippocampal formation is intact in transverse hippocampal slices, the use of somewhat thicker slices (600 km compared with 400-500 pm as normally used) that contain more cells and a larger connectivity of cells may facilitate the detection of synchronous synaptic potentials under physiological conditions in the slice (Schwartzkroin and Haglund, 1986). I3. DISINHIBITION Ample experimental evidence supports the notion that impairment of synaptic inhibition plays a pivotal role in the generation and development of epileptiform activity in many models of epilepsy (Dingledine and Gjerstad, 1980; Schwartzkroin and Prince, 1980a; Wong and Prince, 1979; Wheal et a/., 1984; Stelzer et a)., 1987). Despite a number of experimental data showing impairments of the inhibitory efficacy, direct proof for a causal link between reduced efficacy of synaptic inhibition and epilepsy in humans is not established. In ziitro, synchronized activity can be modified by pharmacological suppression of GABA,4 receptor sensitivity (Fig. 18) (Miles and Wong, 1987a; cf. Traub ~t al., 1989a,b). Using concentrations somewhat lower than those required to block GABA, receptors completely or during the wash-in of GABA, antagonists (“tuning”), the development of synchronization and distinct levels of the synchronization state of a neuronal population can be studied (cf. Traub et al., 1989b): “fast” synaptic inhibition (GABA, mediated) effectively limits the development and propagation of synchronized synaptic activity through a variety of the following mechanisms. 1. Suppresston of Bursts

Although burst firing of CA1 dendrites and CA3 pyramidal cells represents an intrinsic property of these neurons (Kandel and Spencer, 1961; Wong and Prince, 1978, 1979) and the depolarizing wave underlying the burst is triggered by intrinsic fast N a + and is maintained by slow Ca2+ conductances (Wong and Prince, 1978), shunting of GABA,-mediated IPSPs effectively limits the duration and size of Cay+-mediated afterdepolarizations, thus shaping the physiological EPSP-IPSP se-

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quence recorded in CAI and CA3 neurons upon orthodromic stimulation of afferent fibers. Orthodromic stimulation in the presence of GABA, antagonists and the elimination of inhibitory shunting reveals the intrinsic burst capacity in CA1 and CA3 neurons (Wong and Prince, 1979; Wong et al., 1979; Schwartzkroin and Prince, 1980a; Alger and Nicoll, 198213). 2. Suppression of Recurrent Excitation As discussed in more detail in Section IV,A (cf. Miles and Wong, 1987a; Traub et al., 1989a,b), burst propagation within the network (CA3) is facilitated as GABA,-mediated inhibition is reduced. With the activation of latent excitatory connections and inhibition fading, individual synaptic inputs become larger and predominantly excitatory. The intervals between events and the spatial domain of spread of activity increase. Eventually, with inhibition completely blocked, fully synchronized bursts occur, affecting a large population of cells, with interburst intervals of several seconds (Traub et al., 1989b). 3. Inhibitory Entrainment

The rhythmical discharge of spontaneously bursting pacemaker cells can be modulated by IPSPs (Perkel et al., 1964; Lopes da Silva et al., 1976; Pinsker, 1977; Freeman, 1979; Pedley et al., 1980); the time course of recurrent inhibition can filter out a particular population frequency. A model of inhibitory entrainment has been described in Aplysiu neurons in which a single inhibitory interneuron can superimpose a uniform rhythm upon spontaneously bursting cells (Pinsker, 1977); the interburst interval in a given cell is determined by the magnitude of Ca2 -activated K currents mediating large burst afterhyperpolarizations; if an IPSP occurs during the afterhyperpolarization, a superimposed IPSP will delay the onset of the next burst and thus prolong the interburst interval. If the IPSP occurs during the burst, burst duration will be short-ended, less Ca2+ will enter the cell, and the after hyperpolarization is proportionally smaller, thus shortening the interburst interval. +

+

C. SYNCHRONIZATION OF THE INHIBITORY CIRCUIT Experimental data from both slice recordings and computer simulations demonstrate that partially synchronized excitatory bursts under physiological conditions activate inhibitory interneurons, which in turn fire in synchronized bursts, thus limiting the growth and propagation of

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the excitatory activity (Traub et al., 1989b). However, burst firing of inhibitory interneurons is also an intrinsic property of the hippocampal network and is not dependent on the excitatory drive: rhythmic synchronized I PSPs occur spontaneously in hippocampal slices from guinea pigs, monkeys, and humans (Schwartzkroin and Haglund, 1986; Traub et ul., 1989b; Ropert et al., 1990). Bath application of bicuculline or picrotoxin blocks synchronized IPSPs completely, indicating that they are mediated by GABA, receptors. Under physiological conditions (in nonepileptic tissue), synchronized spontaneous IPSPs are specific for the hippocam pal pyramidal cell region, because they are not observed in dentate gyrus o r in neocortical slices (Schwartzkroin and Haglund, 1986). Synchronized IPSPs are triggered by bursts of interneurons that produce an invariably depolarizing synaptic potential in other interneurons and mixed depolarizing/hyperpolarizing postsynaptic potentials in hippocampal pyramidal cells (Schwartzkroin and Haglund, 1986; Muller and Misgeld, 1990). Similar synchronized IPSPs were observed in paired intracellular recordings of CA3 guinea pig hippocampal slices and in simulation studies from pyramidal cells (cf. Traub et al., 1989a). In monkey and guinea pig hippocampal slices, spontaneous IPSPs due to interneuron firing are rhythmic (2-sec intervals) and burstlike reaching peak frequencies of up to 200 Hz during bursts (Ropert et al., 1990). Combined extra- and intracellular recordings and intracellular recordings from pairs of' cells demonstrate a remarkable enhancement of inhibitory synchrony in the presence of 4-AP: with excitatory synaptic transmission blocked, intracellular recordings from neocortex cells show that during type I1 field discharges, all principal cells exhibited synchronous giant IPSPs (Arani et al., 1991). Extracellular field activity in the hilus is time-locked with similar activity in the granule cell layer and in CA3 (Muller and Misgeld, 1990). Paired recordings demonstrate that bursts in hilar interneurons occur synchronously with bursts or giant IPSPs in other hilar interneurons and giant IPSPs in granule cells and pyramidal cells of CA3 and CAI (Muller and Misgeld, 1990; Michelson and Wong, 1991). Although interneuron bursts are more frequently observed in hilar interneurons, the generation of GABA,-mediated bursts in interneurons and giant IPSPs in principal cells and interneurons is not confined to a specific hippocampal subfield, as both events persist after isolation of individual hippocam pal subregions (Michelson and Wong, 1991). Similarly, GABA,-mediated long-lasting depolarizing potentials (LLDs) in the presence of 4-AP appear simultaneously in all areas of the hippocampus (Perreault and Avoli, 1989). After the complete isolation of

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each hippocampal subregion, LLDs can be generated independently in each (isolated) subfield and-similar to interneuron bursting-no specific pacemaker region for LLDs is identified. In nonepileptic tissue of monkeys, synchronous IPSPs were recorded exclusively in interneurons and principal cells of the hippocampal Ammon’s horn, whereas in slices of human epileptic tissue synchronous IPSPs were observed in many cortical areas, neocortical slices, and all hippocampal subfields, including dentate gyrus (Schwartzkroin and Haglund, 1986). In the presence of 4-AP, with glutamate-mediated transmission blocked, synchronized inhibitory activity is also exhibited by the entire hippocampus, including dentate gyrus and neocortex. Although each hippocampal subfield is independently capable of producing inhibitory bursts, hilar interneuron bursting acts as pacemaker for other hippocampal areas in the intact hippocampal slice (Muller and Misgeld, 1990; Michelson and Wong, 1991). With excitatory amino acid-mediated transmission intact, synchronized bursting is a feature of both principal cells and interneurons: however, hilar interneurons burst more easily than the principal cell counterparts (Muller and Misgeld, 1990) and bursts in inhibitory neurons lead burst activity in excitatory pyramidal cells (Schwartzkroin and Haglund, 1986). Synchronized inhibitory activity is of synaptic origin: low extracellular Ca* or removal of extracellular Ca2 blocks inhibitory type I1 responses in neocortex (Aram et al., 1991) and spontaneous rhythmic synchronized activity and long-lasting depolarizations in the hippocampus (Schwartzkroin and Haglund, 1986; Perreault and Avoli, 1989). Bath application of GABA, antagonists bicuculline or picrotoxin blocks all synchronized activity, 4-AP evoked (Muller and Misgeld, 1990; Perreault and Avoli, 1989; Michelson and Wong, 1991) or physiological synchronized activity (Schwartzkroin and Haglund, 1986). Inhibitory synchronized activity is mediated by synchronously firing GABAergic interneurons and remains present in the absence of glutamate-mediated excitation (blocked by CNQX and APV) (Muller and Misgeld, 1990; Michelson and Wong, 1991). The propagation of synchronized inhibitory activity occurs via at least two different transmission processes, one synaptical and one nonsynaptical. This notion is based on two observations. First, in many cases local application of bicuculline does not block rhythmic IPSP activity; the burst generation in specific areas triggers the onset of synchronized inhibitory activity, which propagates to other areas of the hippocampal formation (Schwartzkroin and Haglund, 1986). Second, paired recordings demonstrate that bursts in hilar interneurons occur simultaneously with bursts or giant IPSPs in other hilar interneurons and giant IPSPs in +

+

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granule cells and CA3/CAl pyramidal cells (Muller and Misgeld, 1990; Michelson and Wong, 1991). A similar synchrony is demonstrated in propagation measurements of 4-AP-induced long-lasting depolarizations that occur simultaneously in different hippocampal subfields (Perreault and Avoli, 1989). Bursting of hilar interneurons in the intact transverse slice sets the pace for rhythmic GABAergic synaptic potentials that can be measured synchronously as giant IPSPs or long-lasting depolarizations in other hippocampal subfields (Muller and Misgeld, 1990; Pei-reault and Avoli, 1989; Aram et al., 1991). 'There are two possible nonsynaptic mechanisms that could play a role in the recruitment and the local synchronization of inhibitory neuronal discharge: electric field effects (ephaptic interactions between cells) and electrotonic coupling via gap junctions. However, interneurons are rather sparsely distributed in the hippocampal formation and the neocortex (Ribak P t al., 1978), and electric field effects are therefore unlikely to be effective. Synchronized inhibitory activity (giant IPSPs) recorded in principal cells in the absence of excitatory synaptic transmission is most likely not transmitted through gap junctions between principal cells, because action potential discharges elicited by current injection into one principal neuron did not induce a response in other principal neurons (Miiller and Misgeld, 1990). However, in paired recordings of interneurons, activation of a given neuron by current injection produced a synchronous response in the second interneuron, indicating that interneurons may be electrotonically coupled (Michelson and Wong, 1991). The notion of electrotonic coupling is supported by anatomical data using electron microscopy, demonstrating the existence of gap junctions between interneurons in the dentate gyrus, CA3, and CA1 (Kosaka, 1983a,b).

D.

SYNCHRONIZAI'ION FOLLOWING

TETANIZATION

Tetanization enhances synchronized firing of cells, both in zriuo and in vitro (Buzsaki, 1984b; Slater et al., 1985; Stasheff et al., 1985; Miles and N'ong, 1987b). The development and propagation of synchronized activity following tetanization represents a complex combination of several mechanisms, synaptic and nonsynaptic, with inhibition as a pivotal causal factor. 1. Disinhihitioii

In the CA3 hippocampal subfield, tetanization of afferent fibers mimics the synchronization effects induced by the pharmacological

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blockade of GABAA receptors; activation of recurrent excitatory pathways and growth and propagation of synchronous synaptic potentials are well (inversely) correlated with the efficacy of synaptic inhibition (Miles and Wong, 1987b). In intracellular recordings from CA 1 pyramidal cells, growing synchronization following tetanization of the Schaffer collaterals in stratum radiatum is reflected by an enhanced depolarizing response to orthodromic stimulation. Epileptiform activity in the form of paroxysmal depolarization shifts occurs after IPSPs are reduced to less than 10% of control values, usually after the fourth tetanus has been applied. At this stage of excitability, GABA, receptor sensitivity as assessed by the amplitude of iontophoretic GABA responses is impaired to a similar degree (Stelzer et al., 1987). T h e in vitro studies in the CA3 and CA1 hippocampal subfield demonstrate a parallel development of growth of synchronization of synaptic potentials and impairment of synaptic inhibition and suggest that disinhibition is most likely a causal mechanism of tetanization-induced increase of excitability and generation of highly synchronized population behavior in both hippocampal subfields (Miles and Wong, 1987b; Stelzer et al., 1987).

2. Synchronization of the Inhibitory Circuit cfollowing Tetanization) In nonpyramidal CA1 interneurons, identified by passive cell membrane properties (brief spikes, short time constant, small input resistance, large afterhyperpolarizations following afterpolarizations, lack of inward rectification upon hyperpolarizing current injection), spontaneous giant IPSPs and inhibitory bursts occur synchronously with small field potentials in slices in which the CA2/CA3 pacemaker region for synchronized activity in CAI is removed by dissection (cf. Section V,A; Fig. 7). These data indicate that tetanization produces-in addition to highly synchronized excitatory activity-synchronized activity of the inhibitory circuitry similar to that observed in the presence of 4-AP (see Section IV,B) (Michelson and Wong, 1991; Muller and Misgeld, 1990; Perreault and Avoli, 1989) (for possible propagation mechanism, see below). Tetanization-induced synchronization of the inhibitory circuit in CAI is surprising in two aspects. First, spontaneous giant IPSPs and inhibitory bursts are generated by the CAI circuitry, which is not able to generate substantial spontaneous excitatory synchronized activity. Weak functional connections between CAI pyramidal cells may be the main reason. Second, GABAA-mediated synchronization, producing bursts and giant IPSPs, develop despite a progressive reduction of GABA, receptor sensitivity following tetanization.

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Several factors could promote synchronization of the inhibitory system following tetanization: 1. Bursting of interneurons: Inhibitory interneurons burst more easily than the principal excitatory cells as assessed by comparing 4-APinduced bursting in hilar interneurons and dentate granule cells (Miiller and Misgeld, 1990). 2. Changes of GABA, receptor sensitivity: In nonpyramidal interneurons, the biphasic control response to iontophoretically applied GABA develops into a monophasic depolarizing GABA response, with the depolarizing component greatly enhanced after tetanic stimulation (Fig. 6). In addition, tetanization of afferent fibers results in an increased firing probability of spontaneously active interneurons (interneuron LTP) (cf. Buzsaki and Eidelberg, 1982; cf. Figs. 6 and 7). Increased excitability of interneurons may promote increased release of GABA, a mechanism that-in analogy to 4-AP-may reveal the GABA,-mediated long-lasting depolarizing response. Both mechanisms, enhanced sensitivity of the depolarizing GABA, response and an increased release of GABA, may contribute to GABA acting as excitatory transmitter. 3. Circuit properties: Inhibitory interneurons in CA 1 fire spontaneously in rhythmic bursts (Ropert et al., 1990). T h e relative increase of firing-induced spontaneous IPSPs after tetanization (due to an enhanced presynaptic efficacy of GABAergic inhibition) and decrease in miniature IPSPs after tetanization (due to reduction of GABA, sensitivity) may shift the relative balance toward evoked spontaneous (rhythmic and burstlike) IPS& and reduce the contribution of desynchronizing miniature IPSPs. 4. MPOs: Membrane potential oscillations occur in both principal cells and interneurons at depolarized membrane potentials (Leung and Yim, 1988) (cf. Fig. 8). In interneurons, the gap between resting membrane potential and AP threshold is smaller and a lowering of the threshold for MPOs in interneurons produces higher oscillatory activity at resting potential following tetanization.

E. GENERAL PROPERTIES OF SYNCHRONIZATION 1. Expression of Synchronization T h e genesis and propagation of synchronized activity constitute complex phenomena involving features of single cells, synaptic transmission, and population behavior. Similar to “epilepsy” in humans, which is not a self-defining pathological entity but rather an unspecific response to

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many different affections of the brain, synchronized activity represents an expression of alterations caused by a wide variety of different mechanisms. Yet the expressions of hyperexcitability and synchronization show a considerable similarity for the various models: epileptiform activity measured in brain slice preparations in the form of interictal-like activity (paroxysmal depolarization shifts) and electroencephalographic seizurelike activity resemble the expression of epilepsy in vivo and represent rather stereotype responses to a variety of underlying mechanisms. There are minor differences in the expression of synchronization: for example, in the presence of picrotoxin and after tetanization, a single cell can induce synchronization and spread of firing in a population of cells (Miles and Wong, 1987a,b),whereas synchronization and spread of firing in 4-AP requires the cooperative activity of more than one cell (Perreault, 1990). Major differences in the expression of synchronized activity, both physiological and pathological, reflect differences in the anatomy and connectivity of the circuit (e.g., between CA3 and CA1 hippocampal subfields). Synchronized Excitatory Activity in CA? and CAI. In CA3 pyramidal cells, spontaneous synchronized burst firing is an intrinsic, physiological property (Wong and Prince, 1981) effectively controlled by synaptic inhibition. The CA2/CA3 hippocampal subfieId has been proposed to act as pacemaker region for the propagation of epileptiform activity based on the observation that blockade of GABA, receptors produces interictal spikes in the CA1 subfield of the intact hippocampal slice, which disappear after severing the Schaffer collaterals from CA3 to CA1 (Schwartzkroin and Prince, 1978; Wong and Traub, 1983). Recurrent EPSPs in CA3, which precede the development of fully synchronized activity in CA3 after blockade of GABA, receptors by picrotoxin and following tetanization (Miles and Wong, 1987a,b), trigger delayed EPSPs in CAI (Gjerstad et al., 1981; Wong and Traub, 1983). A similar delayed EPSP in CAI is abolished by focal application of T T X over the CA2/CA3 subregion and occurs in the presence of 4-AP (Perreault and Avoli, 1989). After the establishment of highly synchronized activity, synchronized bursts can be elicited by localized stimuli anywhere in the CA3 region, indicating that the network of excitatory connections in the CA3 region is statistically isotropic (Wong and Traub, 1983). In the isolated CAI hippocampal subfield, the generation of a synchronized response, e.g., in form of a PDS, is pathway specific: repeated tetanization of the Schaffer collaterals results in PDS-like responses upon orthodromic conditioning stimulation whereas antidromic stimulation of alveus/oriens fibers does not produce PDS-like events (unpublished observation). The capability of cells to burst and a strong excitatory connectivity, features characteristic for CA3, seem to be pre-

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requisites for producing spontaneous synchronized excitatory activity. T h e sparsity and weakness of local excitatory connections in CA1 compared with CA3 (recurrent excitation) (Christian and Dudek, 1988) may preclude the genesis of measurable spontaneous excitatory activity in CAI (Brown et al., 1979; Alger and Nicoll, 1980; Ropert et al., 1990). Analogous to the burst propagation within the disinhibited CA3 subfield, stimulation of orthodromic fibers (Schaffer collaterals) produces burst responses in CAI instead of the physiological EPSP-IPSP sequence following GABA,, receptor block (Masukawa and Prince, 1984; Wong ~t al., 1979). In addition, spontaneous epileptiform activity generated in CA3 can propagate from CA3 to the CA1 region regardless of the underlying mechanisms. In summary ( 1) stimulation-evoked excitatory synchronized activity can be generated in the isolated CA1 subfield. The generation of synchronized activity is a genuine property of the CA1 hippocampal subfield and not dependent on input from the CA2-CA3 region, as it is observed in transverse slices in which the CA2-CA3 region has been cut off. <:A 1, like other areas of the hippocampus and neocortex (cf. Traub et al., 1989a), possesses the intrinsic capability for synchronized activity to a certain degree. (2) Epileptiform burst activity in CA1 is contingent upon stimulation of afferent fibers and does not occur spontaneously. It is also pathway specific and confined to the apical dendrite area in stratum radiatum. CA 1 is-in contrast to CA3-statistically not isotrop. In contrast to excitatory synchronized activity, spovitaneous inhibitory synchronized activity is an intrinsic property of the CAI subfield, as it is observed in transverse slices cut off from the CA2-CA3 subfield in the presence of 4-AP (Perreault and ,4voli, 1989; Aram et al., 1991; Muller and Misgeld, 1990) and following tetanization (Fig. 7).

2. Synchronized Inhzbitwn uersw Synchronized Excitation Several pivotal features of synchronized inhibitory activity are in marked contrast to the role of inhibition in the synchronization of excitatory synaptic activity proposed for the CA3 circuitry: in the latter, inhibitor) activity is driven either by afferent synapses (Buzsaki, 1984a) o r by axon collaterals of nearby pyramidal cells. Synchronized bursts originate mainly from excitatory cells and the spread and amount of synchronized activity is effectively controlled by the efficient inhibitory circuit (cf-. Miles. 1990). Synchronization is a function of the reduction of GABA,-mediated inhibitory efficacy (cf. Traub et al., 1989b). T h e strength of excitatory (positively correlated) and inhibitory synapses (inversely correlated) influences the amplitude and frequency of population oscillations (Traub et al., 1989a). Both the origin and the propaga-

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tion are synaptic. Growing synchronization reflects the increase of synchronized synaptic potentials in amplitude and a decrease in frequency (Traub et al., 1989a). Similar mechanisms apply when hyperexcitability and synchronization in the CA3 circuitry are generated by tetanic stimulation of afferent fibers (Miles and Wong, 1987b). Synaptic inhibition in vivo may serve a double function in the control of synchronized activity. Locally, circuit and synaptic properties of inhibition effectively confine the spread and build-up of (excitatory) synchronized activity. GABAA-mediated shunting suppresses late depolarizing synaptic and intrinsic components, giving shape to a physiological EPSP-IPSP sequence upon orthodromic stimulation in the entire CA3 and in CAl dendrites. The efficiency of synaptic inhibition limits the propagation and growth of (excitatory) synchronized bursting (Miles and Wong, 1987a; Traub et al., 1989b). Globally, inhibitory interneurons set the pace and the rhythm for synchronized activity in both inhibitory and excitatory neurons. Giant IPSPs (spontaneous rhythmic synchronous events) in interneurons are depolarizing triggering multiple action potentials. In simultaneous recordings of pyramidal cells and interneurons, the interneuron discharge led the (multiphasic) giant IPSP in pyramidal cells (Schwartzkroin and Haglund, 1986). Such a pacesetting role of inhibitory neurons for a large group of cells, including excitatory cells (“inhibitory phasing”), was proposed in the 1960s by several authors in studies of alpha rhythm generation (Andersen and Sears, 1964; Andersen and Anderson, 1968; Andersen and Eccles, 1962). Tetanization-induced synchronized activity combines features of both aspects of inhibition in regulating and promoting synchronization: (1) reduction of GABA, receptor sensitivity, leading to disinhibition and the growth and spread of (excitatory) synchronized activity (Miles and Wong, 1987b) (much less is known about the role of GABA, receptors, the sensitivity of which is equally reduced after tetanic stimulation); and (2) enhancement of inhibitory synchronized activity. T h e reduction of GABAA inhibition as assessed by the reduction of evoked and spontaneous IPSPs and GABA sensitivity in pyramidal cells is most likely the causal factor in the generation of the (excitatory) synchronized activity observed following tetanization in both CA3 (Miles and Wong, 1987b) and CAI (Stelzer et al., 1987). Tetanization-induced changes of GABA receptor sensitivity in interneurons, in which the hyperpolarizing GABA component fades and the depolarizing component increases (Fig. 6), could contribute to spontaneous giant IPSPs and GABAergic bursts following repeated tetanization. The pivotal factor that determines the degree of synchronized inhibitory activity-as

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inferred from 4-AP experiments-is most likely enhanced release of GABA. Synchronized activity in the presence of 4-AP is not accompanied by changes in the sensitivity to either glutamate, GABA, or NMDA (Perreault, 1990), indicating a presynaptic mechanism for both excitatory and inhibitory synchronized activity. Tetanization increases the presynaptic strength of synaptic inhibition by various means (see above). T h e question as to whether and as to what degree increased interneuron excitability following tetanization results in increased release of GABA remains to be elucidated. Simulation studies show that excitatory synchronization can, in addition to disinhibitory mechanisms and in analogy to synchronization of inhibitory activity, also be achieved by (solely) strengthening the efficacy of excitatory synapses (Traub et d., 1989b). Mechanisms of a possible link between increased release of excitatory o r inhibitory transmitter and synchronization of excitatory and inhibitory activity, respectively, remain to be elucidated. In the isolated CA 1 hippocampal subfieled, tetanization-induced giant IPSPs and GABAergic (PDS-like) bursts are solitary, nonrhythmic, infrequent events, the frequency of which is not dependent on the holding potential. Giant IPSPs and depolarizing bursts were not observed until several tetani had been administered (usually after three to four tetani). Factors such as the general reduction of GABA sensitivity in principal cells and the isolation of the CAI hippocampal subfield, in which synchronization properties are expressed to a lesser extent cornpared with the intact slice, may explain the small extent of synchronization, especially when compared with synchronized activity evoked by bath-applied 4-AP. However, the occurrence of spontaneous IPSPs as such represents a surprising property of the CA1 hippocampal subfield in light of the lack of spontaneous synchronized excitatory activity in <:A I . Tetanization-induced depolarizing GABA potentials in interneurons, both the depolarizing response to iontophoretic GABA and the spontaneous GABAergic burst responses, are accompanied by action potential firing similar to interneuron bursts evoked in 4-AP. Depolarizing GABA., responses in interneurons lack shunting properties and are “excitatory.” ‘This feature is in marked contrast to long-lasting GABA, depolarkations in principal cells in the presence of 4-AP: although depolarizing at physiological resting membrane potentials (reversal at about 5-20 mV above resting potential), LLDs prevent cell firing even when the depolarizing potential is close to o r above firing threshold (Perreault and Avoli, 1989). LLDs act “inhibitory”: their function may represent a specialized form of inhibition, but the exact functional consequence remains elusive. T h e physiological role of synchronized inhibition and the functional

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consequences of enhancement of synchronized inhibitory activity are unknown. The synchronized inhibitory circuit with enhanced inhibitory synaptic potentials, both spontaneous and stimulation evoked, suggests an actual enhancement of the efficacy of synaptic inhibition. But synchronization of inhibitory synaptic activity may effectively reduce the overall efficacy of the inhibitory circuit: (1) Synchronized inhibitory bursts and consecutive massive releases of GABA result in depolarizing GABA responses (cf. Perreault and Avoli, 1988). (2) GABAB-mediated inhibition acts globally by hyperpolarizing the cell membrane without large conductance changes and thus reducing the general excitability of the cell (cf. Section 111,A). Excessive activation of GABA, receptors exerting shunting at a local level may compromise the global inhibitory check by the GABAB system. (3) random distribution of spontaneous IPSPs may be critical in the control of a population’s behavior. Synchronization of inhibition produces a shift from random inhibitory signals toward regular, rhythmic signals. Simulation studies show that steady-state inhibitory inputs produce a more efficient inhibitory control than transient inputs, regardless of specific time courses (Koch, 1985). The (relative) loss of tonic inhibition plays a potentially crucial role in the increase of excitability, although regular IPSPs, both evoked and spontaneous, may not be changed in amplitude or duration or may even be bigger and longer (Aram et al., 1991). Because discharge-evoked spontaneous IPSCs occur in a fairly regular rhythm, whereas miniature IPSCs are random (Ropert et al., 1990), it is conceivable that miniature IPSCs add the “noise” signals to produce a continuous Fourier spectrum for the sum of both spontaneous inhibitory synaptic signals thus creating the “chaotic,” irregular inhibitory input necessary for regular brain activity. T h e ratio of miniature IPSPs to evoked rhythmic sIPSPs could conceivably be shifted by tetanization of afferent fiber pathways by two mechanisms: first, by tetanization-induced synchronization of inhibition; second, it is conceivable that slight decreases in the sensitivity of GABA,-mediated inhibition may selectively eliminate miniature IPSPs, which may functionally result in highly synchronized activity with “inhibition intact” (i.e., evoked inhibitory potentials unchanged or increased).

Acknowledgments

I thank A. R. Kay, R. Miles, and P. Bergold for helpful discussions and R. Miles and R. K. S. Wong for permission to use Fig. 18. Studies were supported by SFB, NIH, Epilepsy Foundation of America, and the Klingenstein Foundation.

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References

Abeles, M. ( 1982). “Local Cortical Circuits.” Springer-Verlag. Berlin. Abraham, W. C . , Gustafsson, B., and Wigstrom, H. (1987).J. Physiol. (London) 394, 367380. Aitken, P. G. (1985). Brain Res. 325, 261-269. Akaike, N., Kaneda, M., Hori, N., and Krishtal, 0. A. (1988). Neurosci. Lett. 87, 75-79. Alger, B. E. (1985). In “Neurotransmitter Action in the Vertebrate Nervous System’’(M. A. Rogawski and J. L. Barker, eds.), pp. 33-69. Plenum, New York. Alger. B. E., and Nicoll, R. A. (1979). Nature (Lotdon) 281, 31.5-317. Alger, B. E., and Nicoll, R. A. (1980). Brain Res. 200, 195-200. Alger, B. E., and Nicoll, R. A. (1982a).J. Physzbl. (Londm) 328, 105-123. Alger, B. E.. and Nicoll, R. A. (1982b).J. Physiol. (London) 328, 125-141. Allen, C . N.,Brady, R., Swann, J., Hori. N., and Carpenter, D. 0. (1988). Brain Res. 458, 147-150. Andersen, P., and Anderson, S. A. (1968). “Physiological Basis of Alpha Rhythm.” Appleton-Century-Crofts, New York. Andersen, P.. and Eccles, J. (1962). Nafure (London) 196, 645-647. Andersen, P., Eccles, J. C., and Loyning, Y. (1963). Nature (London) 198, 540-542. Andersen, P., Eccles, J. C., and Loping, Y. (1964a).J . ,Veurophysiol. 27, 592-607. Andersen, P., Eccles. J. C., and Loyning, Y . (1964h). J . Neurophysiol. 27, 608-619. Andersen, P., and Sears, T. A. (1964).J. Physiol. (London) 173, 499-480. Andersen, P., Dingledine, R., Gjerstad, L., Langmoen, I. A., and Mosfeldt-Laursen, A. M. (1980a).J. Physiol. (London) 305, 279-296. Andersen, P., Sundberg, S. H., Sveen, O., Swann, J. W., and Wigstrom, H. (198Ob). J. Physiol. (London) 302, 463-482. Anderson, C. R.. and Stevens, C. F. (1973).J. Physiol. (Loruton) 235, 655-691. Andrade, R.. Malenka, R. C., and Nicoll, R. A. (1986). Science 234, 1261-1265. Aram, J., Michelson, H.. and Wong, R. K. S. (1991).J. Neurophysiol. 65, 1034-1041. Armstrong, D. L. (1989). Trends Neurosci. 12, 117-122. Armstrong, D. I-., and Eckert, R. (1987). Pror. Natl. Acad. Sci. U.S.A. 84, 2519-2522. k t o k d , A., and Singer, W.91987). Nature (London) 330, 649-652. Ashbv, C. D.. and Walsh, D. A. (1973).J. Biol. Chem. 248, 1255-1261. Awapara, J., Landua, A. J . , Fuerst, R., and Seale, B. (1950). J . Biol. Chem. 187, 35-39. Ayala, G. F., Dichter, M. A., Gumnit, R. J., Matsumoto, H., and Spencer, W. A. (1973). Brain Res. 52, 1-18. Balazs, R.,Hack, N., and Jorgensen, 0. S . (1988). Neurosci. Lett. 87, 80-86. Baraban, J. M., Synder, S. H., and Alger, B. E. (1985).J. Neurosci. 8( 1 I), 4069-4078. Beavo, J . A., and Murnby. M. C. (1982). Handb. Exp. Phurmacol. 58( I), 363-392. Beavo, J. A., Bechtel, P. J., and Krebs, E. G. (1974). Proc. Natl. Acud. Sci. U.S.A. 71, 35803583. Bekkers, J. M., and Stevens, C. F. (1990). Nature (L07tdOlh) 346, 724-729. Berridge, M. J., and Irvine, R. F. (1984). Nature (Lordo7a) 312, 315-321. Berridge, M. J., and Taylor, C. W. (1988). Cold Spring Harbor Symp. Quant. Biol53(2), 927933. Bliss, T. V. P., and Loerno, T. (1973).J. P h y . d . ( h n d O 7 1 ) 232, 331-356. Bliss, T. V. P., and Lynch, M. A. (1988). In “Long-Term Potentiation: From Biophysics to Behavior” (P. W. Landfield and S. A. Deadwyler, eds.), pp. 3-72. Liss, New York. Bloom, F. E., and Iversen, L. L. (1971). Nafure (Lotdon) 229, 628-630.

GABAA RECEPTOR CONTROL

279

Blumenthal, D. K., Takio, K., Hansen, R. S., and Krebs, E. G. (1986).J, B i d . Chem. 261, 8140-8 145. Bormann, J., Sakmann, B., and Seifert, W. (1983).J. Physiol. (London) 341, 9P-10P. Bowery, N. G. (1989). Trends Pharmacol. Sci. 10, 401-407. Brandt, S. J., Nicdel, J. E., Bell, R. M., and Young, W. S., 111 (1988). Cell (Cambridge, Mass.) 49, 57-63. Brown, T. H., Wong, R. K. S., and Prince, D. A. (1979). Brain Res. 177, 194-199. Browning, M. D., Bureau, M., Dudek, E. M., and Olsen, R. W. (1990). Proc. Natl. Acad. Sci. U.S.A. 87, 1315-1318. Buckle, P. J., and Haas, H. L. (1982).J. Physiol. (London) 326, 109-122. Burns, D. B. (1958). “The Mammalian Cortex.” Arnold, London. Buzsaki, G. (1984a). Prog. Neurobiol. 22, 131-151. Buzsaki, G. (1984b). Brain Res. 300, 179-182. Buzsaki, G., and Eidelberg, E. (1982). J. Neurophysiol. 48, 597-607. Byerly, L., and Hagiwara, S. (1988). In “Calcium and Ion Channel Modulation” (A. D. Grinnel, D. Armstrong, and M. B. Jackson, eds.), pp. 3-18. Plenum, New York. Byerly, L., and Moody, W. J. (1984).J. Physiol. (London) 352, 637-652. Byerly, L., and Yazejian (1986).J. Physiol. (London) 370, 631-650. Cassell, M. D., and Brown, M. W. (1977). Lye Sci. 21, 1187-1891. Castagna, M., Takai, Y., Kaibuchi, K., Sano, K., Kikkawa, U., and Nishizuka, Y. (1982).J. Biol. Chem. 257, 7847-785 1. Chad, J. E., and Eckert, R. (1986). J. Physiol. (London) 378, 31-51. Chad, J. E., Kalman, D., and Armstrong, D. (1987). In “Cell Calcium and the Control of Membrane Transport” (L. J. Mandel and D. C. Eaton, eds.), pp. 167-186. Rockefeller Univ. Press, New York. Chen, Q. X., Stelzer, A., Kay, A. R., and Wong, R. K. S. (1990).J. Physiol. (London)420,207221. Chestnut, T. J., and Swann, J. W. (1988). Epilepsy Res. 2, 187-195. Cheung, W. Y. (1980). Science 207, 19-27. Christian, E. P., and Dudek, F. E. (1988).J. Neurophysiol. 59, 90-109. Cohen, P. (1980).In “Molecular Aspects of Cellular Regulation” (P. Cohen, ed.), Vol. 1, pp. 1-255. ElsevierlNorth-Holland, Amsterdam. Cohen, P. (1989). Annu. Rev. Biochem. 58, 453-508. Cole, A. J., Saffen, D. W., Baraban, J. M., and Worley, P. F. (1989). Nature (London) 340, 474-476. Collingridge, G. L., and Bliss, T. V. P. (1987). Trends Neurosci. 10, 288-293. Collingridge, G. L., Kehl, S. J., and McLennan, H. (1983).J. Physiol. (London) 334, 19-31. Collingridge, G. L., Gage, P. W., and Robertson, B. (1984).J. Physiol. (London)-356, 551564. Connor, J. A,, Wadman, W. J., Hockberger, P. E., and Wong, R. K. S. (1988). Science 240, 649-653. Cotman, C. W., and Iversen, L. L. (1987). Trends Neurosci. 10, 263-265. Cull-Candy, S. G. (1984). Proc. R. SOC.London, Ser. B . 221, 375. Curtis, D. R., and Johnston, G. A. R. (1970). In “Handbook of Neurochemistry” (A. Lajtha et al., eds.), Vol. 3, pp. 115-134. Plenum, New York. Curtis, D. R., Felix, D., and McLennan, H. (1970). Br.J. Phannacol. 40, 881-883. Curtis, D. R., Duggan, A. W., Felix, D., Johnston, G. A. R., and McLennan, H. (1971). Bruin Res. 33, 57-73. Demaille, J. G., Peters, K. A,, and Fischer, E. H. (1977). Biochemistry 16, 3080-3086. DeRiemer, S. A., Strong, J. A., Albert, K. A., Greengard, P., and Kaczmarek, L. K. (1985). Nature (London) 313, 313-315.

280

AKMIN STELLER

Dichter, M., and Spencer, W. A. (1969). J. Neurophysiol. 32, 663-687. Dingledine, R., ed. (1984). “Brain Slices.” Plenum, New York. Dingledine, R., and Gjerstad, L. (1980). J. Physiol. (London) 305, 297-313. Dingledine, K..and Langnioen, I. A. (1980). Brain Res. 185, 277-280. lljorup, A., Jahnsen, H., and Mosfeldt-Laursen, A. (1981).Brain Res. 219, 196-201. Doerner. D., Pitler, T. A . , and Alger, B. E. (1988).J. Neurosci. 8( 1 I), 4069-4078. Doroshenko, P. A,. Kostyuk, P. G., Martynyuk, A. E., Kursky, M. D., a n d Vorobetz, 2. D. (1984). .Veurosrwrice 7, 263-267. Douglas, R. M., Goddard, G. V., and Riives, bf. (1982). Brain Res. 240, 259-272. Dunwiddie. T.. and Lynch, G. (1978).J. PltysioL (London) 276, 353-367. Dutar, P., and Nicoli, R. A. (1988). A‘afure (London) 332, 156-158. Eccles, J. (1933). “ l h e Neurophysiological Basis of Mind: T h e Principles of Neurophysiology.” Oxford Univ. Press (Clarendon), London and New York. Eckert, R., and Chad, J. E. (1984). Prog. BzopItys. Mol. B i d . 44,215-267. Eckstein, F. (198.5). Artnu. Herr. Biochem. 54, 367-402. Edwards, F. A., and Gage, P. W. (1988). ,Vcurosci. Lett. 84, 266-270. Edwards, F. A,, Konnerth, A,, Sakmann, B., and Takahashi, T. (1989). Eur.J. Physiol. 414, 600-6 12. Enna, S. J.. and Karbon. E. W. (1987). In “Benzodiazepirie/GABA Receptors and Chloride Channels; Structural and Functional Properties” (R. W. Olseri and J. C. Venter, eds.). Liss, New York. Erondu. N. E., and Kennedy, M. B. (1985).J. Neurosci. 5, 3270-3277. Faber, D. S., and Korn, H. (1982).J. iVeurophy&l. 48, 654-678. Farber, J . I.. (1981). Life Sci. 29, 1289-1295. Farber, L. H., Wilson, F. J.. and Wolff, D. J. (1987).J. Neurochem. 49, 404-414. Finch, D. M., Nowlin, N. L., a n d Babb, T. L. (1983). Brain Res. 271, 201-216. Fonnum, F. (1968). Biochem.J. 106, 401-412. Fonnuni, F.. and Walberg, F. (1973). Brain Res. 54, 115-127. Freeman, W. J. (1979). Biol. Cybeniet. 35, 21-37. Freund, ?I F., and Antal, M. (1988). Nature (London) 336, 170-173. Friedlander, M.J.. Sayer, R. J., and Redman, S. J. (l990).J.Neurosci. 10, 814-82.5. Frotscher. M., Leranth, C . , Luebbers K., and Oertel, W. H. (1984). Neurosci. Lett. 46, 137143.

27, 101 1-102.5. Fujita, Y., and Sam, I-.(1964)./. Neuropl~~siol. Gage, P. W., and Robertson, B. (1985). BI-.J. P h u m c o l . 85, 675-68 1. Gallindo, A. (1969).Brazit Rps. 14, 763-767. Galvan, M.,Grafe, P.. and ten Bruggencate, G. (1982). Brain Res. 241, 75-86. Gamrani, H., Onteniente, B., Seguela, P., Geffard, ni., and Galas, A. (1986). Brain Res. 364, 30- 38 . Gardner-Medvin, A. R. (1976). Pror. R. Soc. London, Ser. B 194, 375-402. Gean, P.-W., and Sliinnick-(;allagher. P. (1989). Brain Res. 480, 160-169. Gill, G. N..and Garren, L. D. (1970). Bzuchm. Biop/ty?c.Res. Commun. 39, 335-343. Gjerstad, L., .4ndersen, P., Langmoen, I. A,, Lundervald, A., and Hablitz, J. J . (1981). Acin Physiol. S c a d . 113, 24.5-252. Gloor, P. (1984). I n “Electrophysiology of Epilepsy” (P. A. Schwartzkroin and H. V. Wheal, eds.), pp. 107- 136. Academic Press, London. Goh, J . W., and Pennefather, P. S. (1988). Science 244, 980-983. Goto, S., Matsukado, Y., Mihara, Y., Inoue, N., and Miyamoto, E. (1986). Brain Res. 397, 161-172. Gray, R., and Johnston, D. (1985).J. “Veurophysiol. 54, 134-142. Gustafsson, B., and WigstrBin, H. (1988). Trends Neurosci. 11, 156-162.

GABA, RECEPTOR CONTROL

28 1

Gustafsson, B., Huang, Y. Y., and Wigstrom, H. (1988). Neurosci. Lett. 85, 77-81. Gyenes, M., Farrant, M., and Farb, D. H. (1988). Mol. Pharmacol. 34, 719-723. Haas, H. L., and Rose, G. (1982).J. Physiol. (London) 329, 541-552. Hablitz, J. J. (1984).J . Neurophysiol. 51, 1011-1027. Hablitz, J. J., and Lebeda, F. J. (1985). Cell Mol. Neurobiol. 5, 353-371. Halpain, S., and Greengard, P. (1990). Neuron 5,237-246. Halpain, S., Girault, J. A., and Greengard, P. (1990). Nature (London) 343, 369-372. Hamill, 0. P., Marty, A., Neher, E., Sakmann, B., and Sigworth, F. J. (1981). P’uegers Arch. 391,85-100. Hamlyn, L. H. (1963).J. Anat. 97, 189-201. Harootunian, A. T., Kao, J. P. Y., and Tsien, R. Y. (1988). Cold Spring Harbor Symp. Quant. Biol. 53(2), 935-943. Harris, E. W., Ganong, A. H., and Cotman, C. W. (1984). Brain Res. 323, 132-137. Harrison, N. L., and Lambert, N. A. (1989). Neurosci. Lett. 105, 137-142. Hartl, F. T., and Roskoski, R., Jr. (1982). Biochemistry 21, 5175-5183. Hebb, D. 0. (1949). “The Organization of Behaviour.” Wiley, New York. Hemmings, H. C., Greengard, P., Lim Tung, H. Y., and Cohen, P. (1984).Nature (London) 310, 503-505. Henn, F. A., and Hamberger, A. (1971). Proc. Natl. Acud. Sci. U.S.A. 68, 2686-2690. Hescheler, J., Kameyama, M., Trautwein, W., Mieskes, G., and Soling, H.-D. (1987). Eur.1. Biochem. 165, 261-266. Heuschneider, G., and Schwartz, R. D. (1989).Proc. Natl. Acad. Sci. U.S.A. 86,2938-2942. Hockberger, P., Toselli, M., Swandula, D., and Lux, H. D. (1989). Nature (London) 338, 340-342. Holsheimer, J., and Lopes da Silva, F. H. (1989). Brain Res. 77, 69-78. Holz, G. G., Rane, S. G., and Dunlop, K. (1986). Nature (London) 319, 670-672. Hosey, M. M., Borsotto, M., and Lazdunski, M. (1986). Proc. Natl. Acad. Sci. U.S.A. 83, 3733-3737. Huang, K.-P. (1989). Trends Neurosci. 12(1I), 425-432. Huettner, J. E. (1988). Soc. Neurosci. Abstr. 14, 5. Ingebritsen, T. S., and Cohen, P. (1983). Eur.J. Biochem. 132, 255-261. Inoue, M., Oomura, Y., Yakushiji, T., and Akaike, N. (1986). Nature (London) 324, 156158. Iversen, L. L., and Neal, M. J. (1968).J. Neurochem. 15, 609-620. Iversen, L. L., Mitchell, J. F., and Srinivasan, V. (1971).J. Physiol. (London) 212, 519-534. Johnston, D., and Brown, T. H. (1981). Science 211, 294-297. Johnston, D., Hablitz, J. J., and Brown, T. H. (1980). Nature (London) 286, 391-393. Kaibuchi, K., Sano, K., Hoshijima, M., Takai, Y., and Nishizuka, Y. (1982). Cell Calcium 3, 323-335. Kairiss, E., Abraham, W. C., Bilkey, D. K., and Goddard, G. V. (1987). BrainRes. 401, 8794. Kameyama, M., Hescheler, J., Mieskes, G., and Trautwein, W. (1987). Eur. J. Physiol. 407, 46 1-463. Kandel, E. R., and Spencer, W. A. (1961).J. Neurophysiol. 24, 243-259. Katz, B., and Miledi, R. (1970). Nature (London) 226, 962-963. Kay, A. R., and Connor, J. A. (1990).J. Neurosci. Methods 33(1), 77-79. Kay, A. R., and Wong, R. K. S. (1986).J. Neurosci. Methods 16, 227-238. Kellerth, J.-O., and Szumski, A, J. (1966). Acta Physiol. Scand. 66, 146-156. Kelso, S. R., Ganong, A. H., and Brown, T. H. (1986).Proc. Natl. Acud. Sci. U.S.A. 83,53265330. Kennedy, M. B. (1988). Cell (Cambridge, Mass.) 59, 777-787.

282

ARMIN S E L L E R

Kikkawa, C., Takai, Y., hiiirakuchi, R., lnohara, S., and Nishizuka, Y. (1982)./. Biol. Chem. 257, I34 1- 1548. King, (;. L., Kncix. J. J.. and Dingledine, R. (1985). Neuroscience 15, 371-378. Kirkness, E. F., Bovenkerk, C.F., Ueda, T.,a n d Turner, A. J. (1989).BZochetn.J. 259,613616.

Klee, C.B., and Cohen, P. (1988). I n “Calniodulin”(P. Cohen and C. B. Klee, eds.), pp. 225-263. Elsevier. Anisterdam. Klee, C:. B., and Vanaman. T. C.(1982). Adz!. Protein Chem. 35, 213-321. Klee, C;. B.. (:roucl~,‘E H., arid Krinks, M.H. (1979). Proc. Nutl. Acud. Sci. U.S.A.76,6720(i273. Klee, (;. 13.. Draetta, G. F., and Hubbard, hl. J. (1988).Adv. Erizymol. 61, 149-200. Knowles, \V. D., and Schwartzkroin, P. A. (1981)./. iVeuro5ci. 1, 318-322. Knowles, W.D., Schneiderman,J. H., Wheal, H. V., Stafstroni, C. E., and Schwartzkroin, 1.’ A. (1984). Cell. ,Mol. Neiirobiol. 4, 207-230. Ktxh, C . (1985). Proc. R. Soc. Loiidon, Srr. B 225, 365-390. Koch, (:., Poggio. T., and Torre, V. (1982). Phi1o.c. Tram. R. Sor. London, Sur. B . 298, 227264. Kiihler, C.. arid Chari-Palay, V. [ 3982). A’ecirosci. Lett. 34, 259-264. Kosaka, T. (1983a). Bt-ain RK 271, 1.57-161. Kosaka, T. (1983b). Bruin Res. 277, 347-351. Kravitz. E. A., lversen, I,. L., Otsuka, M., and Hall, Z. W. (1968). I n “Structure and Function of Inhibitory Neuronal Mechanisms” (C. \‘on Euler, S. Skoglund, and U. Soderberg. eds.), pp. 37 1-376. Perganion Press, Oxford. Krebs, E. G. (197.5). Curr. T@. Cell. Reg. 5, 99-133. Krebs. E. G., and Beavo, J. A. (1979). AJUZU.Rev. Biorhe?~.48, 923-959. Krinks. M. H., Klee, C. B., Pant, H. C., and Gainer, H . (1988)./. zveur05CI. 8, 2172-2182. Krnjevic, K. (1974). Plqsiof. Rezi. 54, 418-540. Krnjevic, K. ( 1983). I7I “Basic Mechanisms ot Neuronal Hyperexcitability” (P. A. Schwartzkroin and H. V. Wheal, eds.), pp. 249-280. Liss, New York. Krnjevic, K., and Schwartz, S. (1967). E x p Brain Res. 3, 320-336. Kriljevic, K., Morris, M. E., and Kopert. N. (1986). Bruin Res. 374, 1-1 1. Kunkel, D. D., Hendrickson, A. E., Wu, J. Y.,and Schwartzkroin, P. A. (1986)./. Neurosci. 6, 541-.5.52.

Lacaille, J. C., and Schwartzkroin, P. A. (1988a). /. Neurosci. 8[4), 1400-1410. Lacaille, J. C., and Schwartzkroin, P. A. (1988b)./. Neurosci. 8(4), 141 1-1424. Lacaille, J. C.,?dueller, A. L., Kunkel, D. D., and Schwartzkroin, P. A. (1987)./. Neurosci. 7, 1979- 1993.

Lacerda, A., Rampe, D., and Brown, A. $1. (1988). Nufure (London) 335, 249-251. Larson, J., and Lynch, G. (1986). Science 232, 98.5-988. Larson, J., Wong, D., and Lynch, G . (1986). Brain Reg. 368, 347-350. Lee, K. S. (1983).J. lVeurosci. 3, 1369-1372. Leidenheimer, N. J . , Hahner, L. D., Browning, M. D., and Harris, R. A. (1990). Soc. Neuro.yci. Abstr. 16, 162.4. Leurig, L. S.. and Yim, C. Y. (1988). Soc. A’eurosci. Ab.cfr. 18, 51.11. Levy, W. B., and Steward, 0. (1979). Bruin Rex. 175, 233-245. Lii1den.D. J., and Routtenberg, A. (1989). Brain Rm. 14, 279-296. Linden, D.J.. M n g , K. L., Sheu, F.-S., and Routtenberg, A. (1988). Bruin Rex. 458, 142146.

Lopes da Silva, F. H., van Rotterdam, A,, Barts, P., van Heudsen, E., and Burr, W. (1976). PWJg.Brnin Re.<.45, 281-308.

GABA, RECEPTOR CONTROL

283

Lorente de NO, R. (1934).J. Psychol. Neurol. 46, 113-177. Lorente de N6, R. (1949). I n “Physiology of the Nervous System” (J. F. Fulton, ed.), pp. 288-312. Oxford Univ. Press, London and New York. LoTurco, J. J., Mody, I., and Kriegstein, A. (1990). Neurosci. Lett. 114, 265-271. Lynch, G., Gribkoff, V. K., and Daedwyler, S. A. (1976). Nature (London) 263, 151-153. MacDonald, R. L., and Barker, J. L. (1979). Brain Res. 167, 323-336. MacVicar, B. A., and Dudek, F. E. (1980). Brain Res. 184, 220-223. MacVicar, B. A., and Dudek, F. E. (1981). Science 213, 782-785. Majewska, M. D., and Chang, D. M. (1984). Mol. Phurmucol. 25, 352-359. Malenka, R. C., Kauer, J. A,, Perkel, D. J., and Nicoll, R. A. (1989). Trends Neurosci. 12,444450. Malinow, R., and Miller, J. R. (1986). Nature (London) 320, 529-530. Malinow, R.,and Tsien, R. W. (1990). Nuture (London) 346, 177-180. Manzoni, 0.J., Finiels-Marlier, F., Sassetti, I., Bockaert, J., Le Peuch, C., and Sladeczek, F. A. (1 990). Neurosci. Lett. 109, 146- 151. Marr, D. (1971). Philos. Tram. R. SOC.London 262, 24-81. Marty, A. (1989). Trends Neurosci. 12, 420-424. Marty, A,, and Neher, E. (1983). I n “Single-Channel Recording” (B. Sakmann and E. Neher, eds.), pp. 107-122. Plenum, New York. Masukawa, L. M., and Prince, D. A. (1984).J. Neurosci. 4, 217-227. Matsumoto, H., and Ajmone-Marsan, 0. (1964). Exp. Neurol. 9, 286-304. Mattson, M. P., Dou, P., and Kater, S. B. (1988).J. Neurosci. 8, 2087-2100. Mayer, M. L., and Westbrook, G. L. (1987).J. Physiol. (London) 394, 501-527. Mayer, M. L., Westbrook, G. L., and Guthrie, P. B. (1984). Nuture (London) 309,261-263. McBurney, R. N., and Neering, I. R. (1987). Trends Neurosci. 10(4), 164-169. McComb, R. B., Bowers, G. N., and Posen, S. (1979). “Alkaline Phosphatase.” Plenum, New York. McCormick, D. A. (1989).J. Neurophysiol. 62(5), 1018-1027. McDermott, A. B., Mayer, M. L., Westbrook, G. L., Smith, S. J., and Barker, J. L. (1986). Nature (London) 321, 519-522. McNamara, J. O., Byrne, M. C., Dasheiff, R. M., and Fitz, J. G. (1980). Prog. Neurobiol. 15, 139- 159. McNaughton, B. L., Douglas, R. M., and Goddard, G. V. (1978). Bruin Res. 157,277-293. Mehta, A. K., and Ticku, M. K. (1990). SOC.Neurosci. Abstr. 26, 162.1. Michelson, H., and Wong, R. K. S. (1991). Science (in press). Mihaly, A., Bencsik, K., and Solymosi, T. (199O).J. Neural Trumm. 79, 59-67. Miles, R. (1990). J. Physiol. (London) 428, 61-77. Miles, R., and Wong, R. K. S. (1983). Nature (London) 306, 371-373. Miles, R., and Wong, R. K. S. (1984).J. Physiol. (London) 356, 97-113. Miles, R., and Wong, R. K. S. (1986).J. Physiol. (London) 373, 397-418. Miles, R., and Wong, R. K. S. (1987a).J. Physiol. (London) 388, 61 1-629. Miles, R., and Wong, R. K. S. (1987b). Nature (London) 329, 724-726. Miles, R., and Wong, R. K. S. (1989).J. Physiol. (London) 373, 397-418. Misgeld, U., and Frotscher, M. (1986). Neuroscience 19, 193-206. Misgeld, U., and Klee, M. R. (1984). Exp. Bruin Res., Suppl. 9, 325-332. Misgeld, U., Sarvey, J. M., and Klee, M. R. (1979). Exp. Bruin Res. 59, 217-229. Misgeld, U., Deisz, R. A,, Dodt, H. U., and Lux, H. D. (1986). Science 232, 1413-1415. Mitchell, R., and Wilson, L. (1984). Neurochem. Int. 6, 387-392. Morrison, J. H., Benoit, R., Magistretti, P. J., Ling, N., and Bloom, F. E. (1982). Neurosci. Lett. 34, 137-142.

284

ARMIN SI‘ELZER

Muller, W., and hfisgeld, U. (1990).J. Neurophyszol. 64, 46-56. Nelson, S. R., and Foltz, F. M. (1983). Exp. Neurol. 79, 763-772. Nestler, E. J., and Greengard, P. (1984). “Protein Phosphorylation in the Nervous System.’’ Wile) (Interscience), New York. Newberry, N. R., a n d Nicoll, R. A. (1984).J. Physiol. (London) 348, 239-254. Newberry, N. R., and Nicoll, R. A. (1985).J. Physioi. (London) 360, 161-185. Nicoll, R. A,, Eccles, J. C . , and Oshima, T. (1975). Nature (London) 258, 625-627. Nishizuka, Y. (1986). Science 233, 305-312. Nishizuka, Y. (1988). Nature (London) 334, 661-665. Nishizuka, Y. (1989).JAMA Am. Med. A ~ s o c .262, 1826-1833. Obata, K., and Takeda, K. (1969).J. ,Veurochenr. 16, 1043-1047. Obata, I<.. Ito, hl., Ochi, R., and Sam, N. (1967). Exp. Brain Res. 4, 43-57. and Venter, J. C., eds. (1986). “Benzodiazepine/GABA Receptors and ChloOlsen, R. W’., ride Channels: Structural and Functional Properties.” Liss, New York. Otis, T. S., Stale!, K. J., and Mody, I. (1990). Sac. h’eurosci. A6s-slr. 16, 76.4. Pasantes-!dorales. H., and Arzate, M . E. (1981). A‘eurosci. R ~ J6,. 465-474. Pearce, I. A., Cmbray-Deakin, h.1. A,, and Burgoyne, R. D. (1987). FEBS Lett. 223, 143147. Pedley, T. A., Traub. R., and Goldensohn, E. S. (1980).In “Pacemaker Neurons in the Mammalian Brain,” pp. 255-268. Pepper, K., Bradley, R. J., and Dreyer, F. (1982). Phjszol. Km. 62, 1271-1340. Perkel, D. H.. Schulman, J. H., Bullock, T. H., Moore, G. P., and Segundo, J. P. (1964). Scierue 145, 6 1-63, Perreault, P. (1990).Dissertation Thesis, McGill University, Montreal. I’erreault, P., and Avoli, M. (1988). Neuroscz. Letf. 89, 293-298. Perreault, P., and Avoli, M. (1989). J . Neurophy.siol. 61, 953-970. Pinsker, H. M. (1977).J. Neurophysiol. 40, 544-556. Pitler, T. A., and Alger, B. E. (1990).Sac. Neurosci. Absfr. 16, 202.4. Porter, N. &I., ‘Twyman. R. E.,Uhler, M. D., and Macdonald, R. L. (1989). Soc. Neurosci. Ahir. 15, 458.2. Prince, D. A. (1968).Exp. IVeuroI. 21, 307-321. Pritchett. D. B., Sontheimer. H., Shivers, B. D., Ymer, S., Kettenmann, H., Schofield, P. R., Seeburg, P. H. (1989). Nafure (Londorr) 338, 582-585. Racine, R. (1978). Neurosurgq 3, 234-252. Ramon v Cajal, S. (1893). “ T h e Structure of Ammon’s Horn.” Neurologie, Leizig. Ramon y (lajal, S. (191 1). “Histologie d u Systenie Nerveux de I’Homme et des Vertebres.” Maloine, Paris. Rane, S.,and Dunlap, K. (1986). Proc. Natl. Acnd. Sci. U.S.A. 83, 184-188. Regehr, W. G., and Tank, D. W. (1990). Nature (London) 345, 807-810. Regehr, W. G., Connor, J. A., and Tank, D. W’.(1989). Nuture (London) 341, 533-536. Reymann, K. G., Briidenianri, R., Kase, H., a n d Matthies, H. (1987). Bruin H a . 461, 388392 Ribak, C. E.. Vaughn, .J. E., and Saito, K. (1978). Brain Re.\. 140, 315-332. Ribak, C. E., Vaughn, J. E., and Barber, R. P. (1981).J. Hirfoclim. 13, 555-582. Roberts, E. (1986). In “BenzodiazepineiGABA Receptors and Chloride Channels” (R. W. Olsen and J. C . Venter, eds.), pp. 1-39. Liss, New York. Roberts, E., and Frankel, S. (19.50).J. B i d . Chenz. 187, 55-63. Roberts, G. W., Woodhams, P. L., Polak, J. M., and Crow, T. J. (1984). Neuroscience 11,35-77. Roberts, P. j.(1974). Bruin Res. 67, 419-4213, Robison, G. A., Butcher, R. W., qnd Sutherland, E. W. (1971). “Cyclic AMP.” Academic Press, New York.

GABAA RECEPTOR CONTROL

285

Ropert, N., Miles, R., and Korn, H. (199O).J. Physiol. (London) 428, 707-722. Roth, L. B., Mehegan, J. P., Jacobowitz, D. M., Robey, F., and Iadarola, M. J. (1989).J. Neurochem. 52, 2 15-22 1. Rutecki, P. A., Lebeda, F. J., and Johnston, D. (1987).J. Neurophysiol. 57, 1911-1924. Saito, K., Barber, R., Wu, J. Y., Matsuda, J.-Y., Roberts, E., and Vaughn, J. E. (1974). Proc. Natl. Acad. Sci. U.S.A. 71, 269-273. Salganicoff, L., and DeRobertis, E. (1965). J. Neurochem. 12, 287-309. Sastry, B. R., Goh, J. W., and Auyeung, A. (1986). Science 232,988-990. Sayer, R. J., Friedlander, M. J., and Redman, S. J. (1990).J. Neurosci. 10, 826-836. Scharfman, H. E., and Sarvey, J. M. (1985). Brain Res. 331,267-274. Schmechel, D. E., Vickrey, B. G., Fitzpatrick, D., and Elde, R. P. (1984).Neurosci. Lett. 47, 227-232. Schneiderman, J. H. (1986). Brain Res. 398, 231-241. Schofield, P. R., Darlison, M. G., Fujita, N., Burt, D. R., Stephenson, F. A,, Rodriguez, H., Rhee, L. M., Ramachandran, J., Reale, V., Glencorse, T. A., Seeburg, P. H., and Barnard, E. A. (1987). Nature (London) 328, 221-227. Schrier, B. K., and Thompson, E. J. (1974).J. BioZ. C h m . 249, 1769-1780. Schwartz, J. H., and Greenberg, S. M. (1987). Annu. Rev. Neurosci. 10, 459-476. Schwartzkroin, P. A., and Haglund, M. M. (1986). Epilepsia 27(5), 523-533. Schwartzkroin, P. A., and Knowles, W. D. (1984). Science 223, 709-712. Schwartzkroin, P. A,, and Kunkel, D. D. (1985).J. Comp. Neurol. 232, 205-218. Schwartzkroin, P. A., and Mathers, L. H. (1978). Brain Res. 157, 1-10. Schwartzkroin, P. A., and Prince, D. A. (1978). Brain Res. 147, 117-130. Schwartzkroin, P. A., and Prince, D. A. (1980a). Ann. Neurol. 7, 95-107. Schwartzkroin, P. A., and Prince, D. A. (1980b). Brain Res. 183, 61-76. Schwartzkroin, P. A., and Wester, K. (1975). Brain Res. 89, 107-1 19. Segal, M. (1 987). Brain Res. 414, 285-293. Seubert, P., Larson, J., Oliver, M., Jung, M. W., Baudry, M., and Lynch, G. (1988). Brain Res. 460, 189-194. Sigel, E., and Baur, R. (1988). Proc. Natl. Acad. Sci. U.S.A. 86, 2938-2942. Sirnan, R., and Noszek, J. C. (1988). Neuron 1, 279-287. Siman, R., Baudry, M., and Lynch, G. (1987). In “Synaptic Function” (G. M. Edelrnan et al., eds.), pp. 519-548. Wiley, New York. Sladeczek, F., Pin, J. P., Recasens, M., Bockaert, J., and Weiss, S. (1985). Nature (London) 317, 717-719. Slater, N. T., Stelzer, A,, and Galvan, M. (1985). Neurosci. Lett. 60, 25-31. Sloviter, R. S., and Nilaver, G. (1987).J. Comp. Neurol. 256, 42-60. Somogyi, P., Smith, A. D., Nunzi, M. G., Gorio, A., Takagi, H., and Wu, J. Y. (1983).J. Neurosci. 3, 1450-1468. Somogyi, P., Hodgson, A. J., Smith, A. D., Nunzi, M. G., Gorio, A., and Wu, J. Y. (1984).J. Neurosci. 4, 2590-2603. Spencer, W. A,, and Kandel, E. R. (1961). Ex#. Neurol. 4, 149-161. Spyker, D. A., Lynch, C., Shabanowitz, J., and Jinn, J. (1980). Clin. Toxicol. 16, 487-497. Srinivasan, V., Neal, M. J,, and Mitchell, J. F. (1969).J. Neurochem. 16, 1235-1244. Stasheff, S. F., Bragdon, A. C., and Wilson, W. A. (1985). Brain Res. 344, 296-302. Stasheff, S. F., Anderson, W. W., Clark, S., and Wilson, W. A. (1989).Science 245,648-651. Stelzer, A. (1990). I n “Excitatory Amino Acids and Neuronal Plasticity” (Y. Ben-Ari, ed.), pp. 255-263. Plenum, New York. Stelzer, A. (1992). I n “Ion Channels” (T. Narahashi, ed.), Vol. 31. Plenum, New York (in press). Stelzer, A,, and Wong, R. K. S. ( 1 987). SOC.Neurosci. Abstr. 13, 179.7.

286

ARMIN STELZER

Stelzer, A, and Wong, R. K. S. (1989).Soc. Neurmci. Abstr. 14, 369.2. Stelzer, A., Slater, N. T., and ten Bruggencate, G. (1987). Nature (London) 326, 698-701. Stelzer, A,, Kay, A. K.,and Wong, R. K. S . (1988). Science 241, 339-341. Stewart, .A. A , , Ingehritsen. t . S., Manalan, A., Klee, C. B., and Cohen, P. (1982).FEBS Lett. 137, 80-84. Storm-hlathisen, J., Leknes, A. K., Bore, A. T., Vaaland, J. L., Edminson, P., Haug, F. M. S.,and Ottersen, 0. P. (1983). Nafut-e(London) 301, 517-520. Sugiyania. H., Ito, I . , and Hirono. C. (1987). Nature (London) 325, 531-533. Sugiyama, H., Ito. I.. and Watanabe, M. (1989).Neuron 3, 129-132. Swanson, L. W’.,Tevler, T. J., and Thompson. R. F. (1982). ilieurosci. Key. Program Bull. 20, 61 1-679. Sweetnam, P. 31., I.loyi,J., Gallombardo, P., Malison, R.T., Gallager, D. W., Tallman,J. F., antl Nestler, E. J. (1988).J. Stwrochrni. 51, 1274-1284. Szente, M . B., and Baranyi. A. fl987). Br&i Rrs. 413, 368-373. Takeuchi, A. (1978).Itr ”GABA in Nervous System Function” (E. RobeI-ts, T. N. Chase, and D. B. Tower. eds.), pp. 2.5.5-267. Raven Press, New York. Taleb, O., Trtruslard.j., Derneneix, B. A . , Feltz, P., Bossu, J.-L., Dupont, J.-L., and Feltz, A. ( 1987). EUV.J . f‘/&y,\id. 409, 620-63 1. Tallant. E. A , , and (:heung. W. Y. (1986).I n “C:alciuni and Cell Function” (W. Y . Cheung, ect.), Vol. 6. pp. 71-1 12. Academic Press, Orlando, Florida. Tamarnaki, N.,Watanabe, K., and Nqjyo, 1’. (1984). Brain Res. 307, 336-340. Tanaka. T.. Ohniura, T., ydniakado, T., and Hiciaka, H . (1982). Mol. Yharmacol. 22, 408412. T;tpia. R., Pasantes, H.. Ortega, B. G.. and Massieu. G. (1966). Biochern. Phanncol. 15, I83 1 - 184.5. ‘l>pia, K., Sancioval. hl., arid Contreras, P. (1975).J. Neurochem. 24, 1283-1285. I.aube, .J. S.. and Schwartzkroin, P. A. (1987). Brain Rex 419, 32-38. Taube, J. S.. and Schwartzkroin, P. A. (1988).J. Neurosci. 8, 1632-1644. Taylor. C:. P.. and Dudek, F. E. (1984).J. iVeuro[hyio[. 52(1), 143-155. ‘fehrani, 51. H. J.. Hablitz, J . J . ,and Barnes, E. M. (1989). Synapse4, 126-131. Thalniann. K. H., Peck, E. 1.. a n d Ayala, G.F. (1981). Neuro.tci. Lett. 21, 319-324. rhonipson, S. $1.. and Gahwiler, 8. H. (l989a). J. .Veurop/tqsiol. 61(3), 501-51 1. ‘Thompsttn.S. M., a i d (Xihwiler. 8. H . (1989b). J. Neurophy,siol. 61(3),512-523. Thompson, S. Xl., and Gahwiler, t3. H. (1989~). J . h’europltysiol. 61(3), 524-533. ‘Tickti. M. K.( 19x6).171 “Briizodiazepine/(;AB.~Keccprors and Chloride Channels: Structural and Functional Properties (R. W.Olsen and J. C. Venter, eds.), pp. 195-5207, Alan

T. (1978). Proc. K . Soc. Lopidoit Ser. El 202, 409-416. Traub, K. D., and \vOng. K. K. S. (1982). Srimce 216, 74.5-747. Traub, R. D., and Wong, R. K. S. (1983).J. , V t u r o p h ~ ~ i49, ~ l . 442-458. Traub, R. D.. Miles, R.,and Wong, R. K. S. (1987).J. Seurophy.\iol. 58, 739-751. Traub, K. D., Miles. K.,and Wong. R. K. S. (1989a). Scicnre 243, 1319-1325. Traub, R . D., Miles, K., and Wong, K. K. S. (198%). “Lectures o n Mathematics in the Life Sciences,” \‘<)I. 21, pp. 6-85. Am. Math. Soc., Providence, Rhode Island. Tsukahara, N. (1981). Rrri. .Vctm.~cI‘.4, 351-379. Utlenfr-iend, S. (1950).J. Bud. C:hcm. 187, 65-69. Vicini, S., A h , H . , Costa. E.. Mienville, J. M., Santi, hl. R.,and Vacarino, F. M. (1986). f‘ror. .Vat/. d4rc~d.Sti. I!..S.A. 83, 9269-9273. Voskuyl, K. X.. and Albus. H. (19x5). Brain R ~ s342, . .54-66. L’yklick!. I... jr., Kriisek,J.. and Edwards. C. (1988).A’eurr,scz. Lett. 89, 313-318.

GABA, RECEPTOR CONTROL

287

Walsh, D. A., and Ashby, C. D. (1973). Recent Prog. Horn. Res. 29, 329-359. Walsh, D. A., Ashby, C. D., Gonzalez, C., Calkins, D., Fischer, E. H., and Krebs, E. G. (1971).J. Biol. Chem. 246, 1977-1985. Watkins, J. C., and Evans, R. H. (1981). Annu. Rev. Pharmacol. Toxicol. 21, 165-204. Watkins, J. C., Krogsgaard-Larsen, P., and HonorC, T. (1990). Trendr Phamucol. Sci. I1 (l), 25-33. Weiss, D. S., Barnes E. M., and Hablitz, J. J. (1988).J Neurophysiol. 59, 495-513. Westerink, B. H., and de Vries, J. B. (1989a). Naunyn-Schmiedeberg’s Arch. Pharmacol. 339, 603-607. Westerink, B. H., and de Vries, J. B. (1989b). Neurosci. Lett. 99, 197-202. Wheal, H. V., Ashwood, T., and Lancaster, B. (1984). In “Electrophysiology of Epilepsy” (P. A. Schwartzkroin and H. V. Wheal, eds.), pp. 173-200. Academic Press, New York. Wigstrom, H., and Gustafsson, B. (1983). Nature (London) 301,603-604. Wigstrom, H., and Gustafsson, B. (1985). Acta Physiol. Scand. 125, 159-172. Wigstrom, H., Gustafsson, B., Huang, Y. Y., and Abraham, W. C. (1986). Acta Physiol. Scand. 126, 317-319. Wilson, R. C., Levy, W.B., and Steward, 0. (1981).J. Neurophysiol. 46, 339-355. Wise, B. C., Guidotti, A., and Costa, E. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 886-890. Wolf, M., KeVine, H., 111, May, W. S., Jr., Cuatrecasas, P., and Sahyoun, N. (1985).Nature (London) 317, 546-549. Wong, R. K. S., and Prince, D. A. (1978). Brain Res. 159(2), 385-390. Wong, R. K. S., and Prince, D. A. (1979). Science 204, 1228-1231. Wong, R. K. S., and Prince, D. A. (1981). J. Neurophysiol. 45, 86-97. Wong, R. K. S., and Traub, R. D. (1983).J. Neurophysiol. 49,442-458. Wong, R. K. S., and Traub, R. D. (1983). Neurology. Wong, R. K. S., and Watkins, D. J. (1982).J. Neurophysiol. 48, 938-951. Wong, R. K. S., Prince, D. A., and Basbaum, A. I. (1979). Proc. Nutl. Acad. Sci. U.S.A. 76, 986-990. Wong, R. K. S., Traub, R. D., and Miles, R. (1986). Adv. Neurol. 44, 583-592. Wood, J. D., and Peesker, S. J. (1972).J. Neurochem. 19, 1527-1537. Wood, J. D., and Peesker, S. J. (1973).J. Neurochem. 20, 379-387. Wood, J. G., Wallace, R. W., and Cheung, W. Y. (1980). In “Calcium and Cell Function” (W. Y. Cheung, ed.), Vol. 1, pp. 291-303. Academic Press, New York. Worley. P. F., Baraban, J. M., and Snyder, S. H. (1986).J. Neurosci. 6, 199-207. Yamamoto, C., and Chujo,T. (1978). Exp. Neurol. 58, 242-250. Yang, S., Tallant, E. A., and Cheung, W. Y. (1982). Biochem. Biophys. Res. Commun. 106, 14 19- 1425. Zimmer, J., Laurberg, S., and Sunde, N. (1983). In “Neurobiology of the Hippocampus” (W. Seifert, ed.), pp. 39-64. Academic Press, New York.

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CELLULAR AND MOLECULAR PHYSIOLOGY OF ALCOHOL ACTIONS IN THE NERVOUS SYSTEM Forrest F. Weight Laboratory of Molecular and Cellular Neurobiology National Institute on Alcohol Abuse and Alcoholism Rockville, Maryland 20852

I. Introduction 11. Lipid Theory of Alcohol Action 111. Alcohol Effects on Neuronal Firing A. Cerebellum B. Hippocampus C. Locus Coeruleus D. Substantia Nigra and Ventral Tegmental Area E. Raphe Nucleus F. Other CNS Regions G. Commentary IV. Alcohol Effects on Cellular Mechanisms A. Electrical Excitability and Voltage-Gated Ion Channels B. Synaptic Transmission and Transmitter-Gated Ion Channels V. Summary and Conclusions A. Ion Channel Hypothesis of Alcohol Action B. Alcohol Sensitivity of Transmitter-Gated Ion Channels and Behavioral Effects of Alcohol C. Conclusions References

1. Introduction

Although the behavioral effects of alcohol have been recognized since ancient times, only recently have we begun to understand the cellular and molecular mechanisms that underlie these behavioral actions. In the 1890s, it was found that the anesthetic potency of different alcohols correlates with their lipid solubility. This led to the development of what has been called the “lipid theory of alcohol action.” Since the introduction of this idea, it has dominated thinking on the nature of alcohol’s actions in the nervous system. In recent years, studies of alcohol’s actions on various types of cellular and molecular phenomena in the nervous system have revealed selective actions that may explain some of alcohol’s behavioral effects. This article will review the physiological 289 INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 33

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actions of alcohol in the nervous system and discuss the possible cellular and molecular basis of alcohol’s behavioral actions. The review will focus particularly on the acute neuronal actions of alcohol, as detected by neurophysiological techniques. Neurochemical effects of alcohol have been reviewed recently (Deitrich et al., 1989; Tabakofr and Hoffman, 1991) and will not be discussed in detail here.

11. Lipid Theory of Alcohol Action

Almost a century ago, Overton (1896, 1901) and Meyer (1899, 1901) found that the anesthetic potencies of alcohols and anesthetics increase in proportion to their partition between olive oil and water. These observations led to the idea that to exert its action, an alcohol or anesthetic agent must dissolve in the hydrophobic lipid bilayer of the membrane, so that the greater the lipid solubility of the agent, the greater its potency. Alcohols exhibit general anesthetic properties, and, up to a point, the anesthetic potency of alcohols increases as their chain length increases (Cole and Allison, 1930; Rang, 1960); a similar relationship obtains for the intoxicating properties of alcohols (McCreery and Hunt, 1978). There have been several variants of the lipid theory. Meyer (1937) suggested that to obtain anesthesia it is necessary to attain a certain molar concentration of the anesthetic agent in the lipid domain of the cell. Ferguson (1939) modified this concept and proposed that anesthetic potency is correlated with the thermodynamic activity of the agent, viz. the concentration of anesthetic times an activity coefficient. Mullins (1954) advanced the hypothesis that anesthesia occurs when a certain critical volume of the membrane is occupied by the anesthetic agent. This idea was extended by Seeman (1972), who proposed that the presence of the anesthetic agent in the membrane expands the cell membrane; this membrane expansion in turn produces conformational changes that result in anesthesia. T h e essential concept in these proposals is that when the anesthetic dissolves in the lipid bilayer membrane, it alters the properties of the membrane lipids and this in turn leads to the development of the anesthetic state. More recently, the interaction of alcohol with cell membranes has been studied using a number of molecular probes and physical techniques such as electron paramagnetic resonance (EPR) or fluorescence spectroscopy. Chin and Goldstein (1977), using EPR and the molecular probe 5-doxylstearic acid, found that ethanol decreased the order parameter in a concentration-dependent manner in synaptosomal mem-

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branes from mouse brain. This was interpreted as evidence that ethanol increases the mobility of membrane lipids; that is to say, the fluidity of membrane lipids increased. To determine whether the ability of different alcohols to fluidize the membrane is related to their lipid solubility, Lyon et al. (1981) studied the relationship of these parameters for a series of alcohols. They found that the membrane-disordering potency of the alcohols correlated with both the membranelbuffer partition coefficient and the hypnotic potency of the alcohols. Such observations led to the hypothesis that the intoxicating and anesthetic properties of alcohols result from their lipid-fluidizing actions in membranes (cf. Goldstein, 1984). In recent years, several criticisms of the lipid theory of alcohol and anesthetic action have been advanced. First, using different membrane probes and different lipid bilayer membranes, several studies have been unable to detect significant effects of alcohols and anesthetics at pharmacologically relevant concentrations (Boggs et al., 1976; Franks and Lieb, 1978, 1979; Turner and Oldfield, 1979; Lieb et al., 1982). Second, when techniques are used that can detect a fluidity increase with pharmacological concentrations of alcohols and anesthetics, the magnitude of the change is extremely small-less than the change of fluidity produced by a temperature increase of 1°C (Franks and Lieb, 1987). It goes without saying that a temperature increase of 1°C does not induce intoxication o r anesthesia in whole animals. Third, the activity of a pure, soluble protein, firefly luciferase, can be inhibited by a number of alcohols and anesthetics, and the potency for inhibiting this protein is correlated with the anesthetic potency of the agents (Franks and Lieb, 1984, 1985). Fourth, the general anesthetic potency of a homologous series of nalcohols falls off when the number of carbons in the chain exceeds 10, in a manner similar to that observed for the inhibition of luciferase, although the partitioning of higher n-alcohols into lipid bilayers does not exhibit such a fall off (Franks and Lieb, 1985, 1986). Fifth, the membrane-fluidizing agent, A,C, produces neither intoxication nor anesthesia (Buck et al., 1989). As noted above, the lipid theory proposes that alcohols exert their primary effects by dissolving in the lipid portions of neuronal membranes and altering their properties. Although the mechanism by which the lipid properties of the membrane are affected varies in different versions of the theory, in general it is suggested that the perturbation of membrane lipids by alcohol and anesthetics secondarily affects the function of membrane proteins involved in the regulation of nerve cell activity. On the other hand, the preceding criticisms of the lipid theory and the observation that pharmacological concentrations of alcohols and

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anesthetics can directly affect the function of a pure protein raise the question of whether alcohols and anesthetics might exert their cellular effects by acting directly on membrane proteins. In view of these considerations, it is important to know what neuronal membrane proteins are particularly sensitive to alcohol, because many proteins have been found to be relatively insensitive (cf. Franks and Lieb, 1982), and to characterize the molecular mechanisms of those actions. In this context, we will now turn our attention to the cellular and molecular actions of alcohol in the nervous system.

111. Alcohol Effects on Neuronal Firing

Because neurons code and conduct information as electrical impulses, one approach to elucidating the cellular actions of ethanol is to study its effects on the spike-firing activity of neurons. A number of studies have reported the effects of ethanol on the frequency and pattern of neuronal spike firing in different brain areas. Those studies will be reviewed briefly below.

A. CEREBELLUM There have been a number of studies of the effects of ethanol on spike firing in cerebellar neurons. This no doubt is due in large part to the fact that the cerebellum plays an iniportant role in motor coordination and control, and ethanol is known to affect these behavioral activities. ‘The neuron investigated in most of these studies is the Purkinje cell, the output neuron of the cerebellar cortex. In early studies on the eKects of ethanol on cerebellar Purkinje cells, Eidelberg et al. (197 1) found, using decerebrate, unanesthetized cats, that the intravenous administration of 0.6 g/kg ethanol decreased the firing rate of these neurons. To test whether this alteration of firing rate might be secondary to ethanol effects in other brain regions, Forney and Klemm (1976) isolated the cerebellum by electrolytic lesions of the cerebellar peduncles. ‘They found, in unanesthetized, curarized rats, that 1 g/kg ethanol injected intraperitoneally depressed impulse activity of most neurons in the cortex of the deafferented cerebellum, and concluded that the effect of ethanol was directly on the cerebellum. Mikesa and Klemm (1979) used D-penicillamine to inhibit the metabolism of

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ethanol to acetaldehyde and reported that the response of Purkinje cells to ethanol administration was not altered by this treatment, leading them to conclude that the effect does not result from a metabolite of ethanol. In subsequent studies, more complex effects of ethanol on the firing of cerebellar Purkinje cells have been observed. Rogers et al. (1980), studying halothane-anesthetized rats, found that 1-4 g/kg intraperitoneal ethanol decreased the mean interspike interval for simple spike firing of Purkinje neurons, but increased the frequency of climbing-fiber bursts in a dose-dependent manner. Using unanesthetized, curarized rats and ethanol doses ranging from 0.25 to 3 g/kg, Chu (1983) found that low doses increased and high doses decreased the spontaneous firing rate of most Purkinje cells; the high doses also increased the frequency of climbing-fiber bursts. In urethane-anesthetized rats, Sorensen et al. (1981a) showed that the intraperitoneal injection of 1-4 g/kg ethanol caused a dose-dependent decrease in the spontaneous discharge of Purkinje cells 30-60 min following administration. In addition, approximately 12 min following ethanol administration, a transient increase in firing rate was observed that returned to control by 30 min. This initial excitation was not observed if the animals were pretreated with 6-hydroxydopamine (to deplete cerebellar norepinephrine) or with the padrenergic antagonist propranolol, suggesting that the excitation is due to a decrease in the tonic inhibitory activity of noradrenergic neurons in the locus coeruleus (see below). Sinclair et al. (1980) and Sinclair and Lo (1981) found that the intravenous administration of 1.5 g/kg ethanol to urethane-anesthetized rats increased the spontaneous spike-firing activity of Purkinje neurons but inhibited climbing-fiber bursts. Climbingfiber bursts result from activity of the climbing-fiber pathway from inferior olivary neurons to Purkinje cells. In rats anesthetized with halothane and in unanesthetized, curarized rats, Rogers et al. (1986) found that the parenteral administration of ethanol increased the firing of inferior olivary neurons, which would increase climbing-fiber bursts in Purkinje cells. On the other hand, Harris and Sinclair (1984b) and Rogers et al. (1986) showed that the parenteral administration of ethanol in urethane-anesthetized rats decreased the firing of inferior olivary neurons, which would explain the reduction in climbing-fiber bursts observed in Purkinje cells in animals anesthetized with this anesthetic. The preceding studies suggest that in in v i m experiments with ethanol administered parenterally, a number of factors can influence ethanolinduced alterations of firing rate; these include the type of anesthetic used, the method and rate of ethanol administration, the concentration

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of ethanol administered, and the time following ethanol administration. These studies also suggest that alterations of firing of one type of neuron may be secondary to ethanol effects on other neurons. To avoid some of these problems, ethanol has also been applied locally to Purkinje cells from micropipettes. Using iontophoresis and micropressure to eject ethanol, Siggins and French (1979) found a concentration-dependent reduction of Purkinje cell discharge. However, the micropipettes contained l or 2 M ethanol and the inhibition of firing was frequently associated with a reduction of spike height and an increase of spike width. In such experiments, the concentration of ethanol affecting the neuron under study is not known; however, the reduction of spike height by ethanol suggests it may have been quite high (see below). Johnson et al. (1985) also found a concentration-dependent decrease in Purkinje cell discharge in response to micropressure application of ethanol; in this case the concentration of ethanol in the pipette was 750 mM and alterations in the extracellular spike configuration were not commented upon. Using in vitro preparations such as brain slices and neurons in tissue culture, ethanol can be applied in known concentrations and a number of indirect effects of ethanol can be obviated. In a study of in uitro cerebellar slices from Simonsen rats, George and Chu (1984) found that the predominant effect of ethanol by bath application was a concentrationdependent reduction of Purkinje cell discharge. In some steady-firing neurons, ethanol concentrations from 9 to 17 mM increased the frequency of firing, but higher concentrations elicited only a decrease of firing rate. On the other hand, in neurons that exhibited a phasic cycling pattern of firing, the lowest concentration of ethanol studied, 9 mM, reduced the firing rate in all neurons tested. In tissue culture experiments using neurons from newborn mouse cerebellum, Seil et al. (1977) showed that bath application of 109 mM ethanol increased both the regularity of firing and the firing frequency. With ethanol concentrations of 370 mM or more, a decrease in firing rate was observed; this was at times preceded by a brief excitation. Franklin and Gruol(l987) studied the sensitivity to ethanol of neurons cultured from fetal rat cerebellum and found that ethanol concentrations as low as 22 mM could alter spontaneous activity of the neurons. Ethanol concentrations from 20 to 80 mM increased the regularity of spontaneous spike firing and either transiently increased or did not change firing rate. The possibility that genetic factors might contribute to the sensitivity of Purkinje cell firing to the administration of ethanol has been tested by studying strains of mice and rats with different behavioral sensitivities to ethanol. T h e inhibition of Purkinje cell firing by pressure ejection of

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ethanol from micropipettes was found to be about twice as sensitive in long-sleep (LS) mice (selectively bred mice that sleep longer than other breeds after a given sedative dose of ethanol) compared to short-sleep (SS) mice (Sorenson et al., 1980; Spuhler et al., 1982). Similarly, an injection of 3.5 g/kg ethanol caused Fisher 344 rats to sleep about 2.5 times longer than Norway Brown rats, and Purkinje cell inhibition by ethanol was found to have about a twofold lower ED,, in Fisher 344 rats compared to Norway Brown rats (Johnson et al., 1985). A differential ethanol sensitivity of Purkinje cell firing between LS and SS mice was also found in cerebellar tissue from fetal LS and SS mice transplanted into the anterior chamber of the eye (Palmer et al., 1982), and in cerebellar slices from LS and SS mice studied in vitro (Basile et al., 1983). The inhibition of cerebellar Purkinje neuron discharge has also been correlated with the loss of righting response in six inbred strains of rat (Palmer at al., 1987). These observations indicate that genetic factors can influence the sensitivity of neuronal discharge to ethanol.

B. HIPPOCAMPUS T h e hippocampus is a brain region of archicortex that is involved in integrative brain functions and certain aspects of learning and memory. T h e structure is particularly well suited for neurophysiological investigation because of the layered organization of cells and defined synaptic pathways. In recent years, some of the cellular and molecular mechanisms involved in learning and memory have been extensively investigated in the hippocampus. Because ethanol is well known to impair cognitive function and the acquisition of information, the hippocampus is a brain region well suited for studying the cellular actions of ethanol. There have been a number of studies of the effects of ethanol on hippocampal neurons; most have focused on pyramidal neurons, the major output cell of the hippocampus. In single unit studies in the hippocampus in vivo using unanesthetized rats, Grupp and Perlanski (1979) found that the intraperitoneal administration of 0.5-3.9 g/kg ethanol produced a concentration-dependent decrease in the frequency of firing of most dorsal hippocampal neurons (predominantly the CA 1 region). At low blood ethanol concentrations (10-60 mg%), an increased firing rate was observed in some neurons. In a subsequent study using a similar preparation, Grupp (1980) observed a biphasic response after intravenous administration of 0.1-0.8 g/kg ethanol. With the lowest dose, the response was an increase in the firing rate o r an excitation followed by some reduction in firing frequency. At

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progressively higher doses, the initial excitation diminished and the subsequent inhibition was augmented, with inhibition of firing observed only at the highest dose of ethanol studied. The effect of local application of ethanol from micropipettes by pressure ejection on the firing of hippocampal CA1 pyramidal cells was studied by Sorensen et al. (1981b) in urethane-anesthetized LS and SS mice. Ethanol decreased the spontaneous firing rate of the neurons in both LS and SS mice; comparison of the sensitivity of the neurons to ethanol in the two types of mice revealed no significant difference. The concentration of ethanol in the micropipette was 750 mM, but, as noted above, the concentration that affects the cell is not known. Berger et al. (1982) applied ethanol from micropipettes containing 1-3 M ethanol using electroosmosis and micropressure ejection in halothaneanesthetized rats. They found that ethanol produced an increased firing rate in 55% of the neurons, a decreased firing frequency in 32% of the cells, biphasic effects (excitation followed by inhibition) in lo%, and no etl'ect in 3%. Because they studied pyramidal neurons in both CA1 and CA3 regions of hippocampus, it is possible that differential sensitivity of neurons in these two regions may contribute to the mixed results (see below). The effect of ethanol on the discharge of pyramidal neurons has also been studied in vitro in the hippocampal slice. Carlen et al. (1982) recorded intracellularly from CA 1 pyramidal neurons and applied ethanol in drops to hippocampal slices on the surface of the bathing solution (interface chamber). They found that ethanol decreased the frequency of spontaneous spike firing in 10 of 12 spontaneously active cells. T h e effect of ethanol was similar at all concentrations of ethanol used (5-20 mM). Using intracellular recordings in submerged hippocampal slices and applying ethanol by bath application, Siggins et al., (1987) found that of' spontaneously firing CA1 pyramidal neurons, 10-350 mM ethanol decreased the firing rate in 5076, had no effect in 29%, biphasically increased and then decreased the firing in 12%, and increased firing frequency in 9%. T h e increased frequency and biphasic responses to ethanol were more common at low (10-43 mM) concentrations of ethanol. Benson et al. (1989) also studied the effect of ethanol on CAl pyramidal neurons recorded intracellularly in hippocampal slice. They found that 50- 100 mh4 ethanol consistently suppressed neuronal firing, with fewer spikes elicited by a 30-sec constant-current pulse in the presence of ethanol. T h e effect was not felt to be attributable to changes of either membrane potential o r membrane conductance. The in uitro hippocampal slice has also been used to study the effects of ethanol on neurons in the CA3 region. Zbicz and Weight (1982) found

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that superfusion of hippocampal slices with ethanol concentrations from 30 to 200 mM produced a concentration-dependent increase in the spontaneous firing of CA3 pyramidal neurons. Intracellular recording revealed that the effect of 30-200 mM ethanol was associated with a decrease in the minimum current needed to initiate action potential generation and an increase in the number of spikes elicited by a constant-current step, viz. a lowering of the threshold for spike initiation and an increase in membrane excitability. No alteration of currentvoltage relationships (I-V curves) was detected at potentials negative to threshold. The results indicate that ethanol in concentrations greater than 30 mM alters membrane excitability of CA3 neurons without an apparent change in membrane conductance. Siggins et al. (1987) also found that CA3 neurons in hippocampal slices were more likely to show excitation in response to ethanol. In their experiments, bath application of 50-200 mM ethanol increased the spontaneous firing frequency of 32% of the neurons, increased and then decreased firing in 32%, decreased firing in 24%,and had no effect in 12%. The administration of ethanol usually was not associated with a change of membrane conductance.

C. Locus COERULEUS Noradrenergic mechanisms have been implicated in alcohol intoxication (Borg et al., 1983) and withdrawal (Linnoila et al., 1987). Norepinephrine-containing neurons are localized, in part, in the nucleus locus coeruleus; these neurons send ascending projections to a number of brain regions, including cortical areas, the limbic system, and the hypothalamus. The norepinephrine released from the terminals of these neurons is thought to modulate the activity of neurons in these regions of the central nervous system (CNS). The effect of ethanol on the activity of noradrenergic neurons in locus coeruleus has been studied by Pohorecky and Brick (1977) in ’ unanesthetized, curarized rats. They found that the intraperitoneal administration of 2 g/kg ethanol decreased the spontaneous firing rate of 62% of the cells tested. The firing rate of these cells was reduced by as much as 80%. In addition, ethanol administration increased the firing frequency of 22% of the neurons tested, and no effect was observed in 16% of the neurons. In rats anesthetized with halothane or chloral hydrate, however, Aston-Jones et al. (1982) did not find any effect of the intraperitoneal administration of 0.5-3 g/kg ethanol on the spontaneous firing rate of locus coeruleus neurons. Similarly, Svensson and Engberg

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(1980) did not find an alteration of firing rate of locus coeruleus neurons after the intraperitoneal administration of 2 g/kg ethanol in chloral hydrate-anesthetized rats. On the other hand, in another study using rats anesthetized with chloral hydrate, Strahlendorf and Strahlendorf (1983) iound that the intraperitoneal injection of 1-2 g/kg ethanol decreased the spontaneous firing frequency in three of four locus COeruleus neurons tested, and the local administration of ethanol to locus coeruleus neurons using electroosmosis inhibited the firing rate in 96% of the neurons. In a recent study of chloral hydrate-anesthetized rats, Verbanck et nl. (1990) reported that the intravenous administration of' 2 g/kg ethanol decreased the firing rate of almost all locus coeruleus neurons studied. I n 60% of the cells there was a very transient excitation preceding the inhibition. The sensitivity of locus coeruleus neurons to ethanol has also been studied in iiitro in brain slices. Shefner and Tabakoff (1985) found that low concentrations ( 1- 10 mM) of bath-applied ethanol reduced the firing rate of some cells, but had no effect or increased firing frequency in decreased other neurons, whereas higher concentrations (30-200 d) the firing rate o f most locus coeruleus neurons. The percent inhibition of' firing f'requency was found to be concentration dependent. The effect of ethanol on locus coeruleus neurons in brain slices has also been investigated by Verbanck et al. (1990). In their study, bath application of ethanol produced significant inhibition of firing at a concentration of 10 miM. A concentration of 100 mM, however, excited about half of the neurons and inhibited the other half'.

NICRAA N D VENTRALTECMENTAL AREA D. SC'BSTANTIA

Dopaminergic mechanisms are thought to be involved in certain types of reward mechanisms (Phillips and Fibiger, 1978; Fibiger et al., 1986), and ethanol has been reported to increase the release of' dopamine in nucleus accunibens (Di Chiara and Imperato, 1988; Wozniak et al., 1990). In addition. destruction of dopamine neurons by 6-OH-dopamine has been reported to decrease alcohol drinking in rats (Meyers and Melchior, 1975). In the brain, dopamine-containing neurons are located primarily in the pars compacta region of substantia nigra and the ventral tegmental area. The axons of pars compacta neurons project predominantly to the neostriatum and central amygdaloid nucleus, and the ventral tegmental neurons innervate nucleus accumbens and the olfactory tubercles.

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1. Substantia Nigra T h e effect of ethanol on the firing of dopaminergic neurons in the pars compacta region (A9) of the substantia nigra has been studied in vivo by Mereu et al. (1984). In unanesthetized, paralyzed rats they found that the intravenous administration of 0.5-2.0 g/kg ethanol increased the spontaneous firing frequency of pars compacta neurons. Increasing the dose increased the duration of the excitatory effect, and doses of 1 and 2 g/kg excited all pars compacta neurons tested. In addition, the firing changed during the ethanol-induced excitation from a regular pattern with fairly uniform interspike intervals to a pattern with bursts of high-frequency spike trains. With cumulative doses of 4 glkg or greater, a reduction of firing frequency was observed after an initial excitation. On the other hand, they found that in rats anesthetized with chloral hydrate or halothane, 1 g/kg ethanol did not affect firing frequency and doses of 2-4 g/kg reduced or completely blocked spontaneous firing. Ethanol-induced excitation of neuronal firing was not observed in the anesthetized preparations. In addition, in unanesthetized preparations, the administration of chloral hydrate, halothane, or pentobarbital increased the firing frequency bf pars compacta neurons. T h e effect of ethanol on the firing of nondopaminergic neurons in the pars reticulata region of substantia nigra was studied by Mereu and Gessa (1985). In unanesthetized, curarized rats they found that the intravenous administration of 0.25-2.0 g/kg ethanol produced a dosedependent reduction in firing frequency in most pars reticulata neurons. T h e effect was rapid in onset and lasted from 15 to 120 min. Of the neurons, 17% did not respond to the doses of ethanol tested.

2. Ventral Tegmental Area The effect of ethanol on the firing pattern of dopaminergic neurons in the ventral tegmental area (A 10) has been investigated by Gessa et al. (1985). They found in unanesthetized, curarized rats that the intravenous administration of 0.125-0.5 glkg ethanol produced dosedependent increases in the firing frequency of ventral tegmental neurons. By contrast, dopaminergic neurons in substantia nigra pars compacts (A9) required ethanol doses of 0.5-2.0 g/kg to elicit increases in firing equivalent to those in ventral tegmental neurons. The ED,, for the firing-rate increase was 0.16 g/kg for ventral tegmental neurons and 0.86 g/kg for nigral pars compacta neurons, indicating that the ventral tegmental dopaminergic neurons were about fivefold more sensitive to the effect of intravenously administered ethanol than were nigral dopa-

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minergic neurons. In rats anesthetized with chloral hydrate, Gessa et al. (1985) did not find effects of ethanol, in doses up to 1 g/kg, on the firing rate of ventral tegmental neurons. On the other hand, Verbanck et al. (1 990), using chloral hydrate-anesthetized rats, found that intravenous (i.v.) administration of 2 g/kg ethanol excited 80%of the ventral tegmental neurons and had no effect or slightly inhibited 20%. T h e effect of ethanol on dopaminergic ventral tegmental neurons has also been studied in uitro in brain slices. Brodie et al. (1990) have reported that bath application of 20-320 mh4 ethanol produced a concentrdtion-dependent increase in the firing frequency of most ventral tegmental neurons studied. The excitatory effect was reversible upon washout of ethanol. In addition, the excitation was observed in a low-Ca2 (0.25 mM)/high-Mg2 (2-8 mM) extracellular medium, suggesting a direct effect of ethanol on the neurons. Verbanck et al. (1990) also found that bath-applied ethanol in concentrations between 10 and 500 mM resulted in a concentration-dependent excitation of ventral tegmental neurons in brain slices. +

+

E. RAPHENUCLEUS Serotonergic mechanisms have been suggested to be important in ingestion and/or craving mechanisms for alcohol. Alcohol consumption in the rat has been reported to be decreased following the intraventricular administration of serotonin (Hill, 1974) or its precursor 5-hydroxytryptophane (Geller, 1973; Meyers and Martin, 1973). In addition, serotonin reuptake inhibitors have been reported to diminish alcohol consumption (Gill and Amit, 1989; Nardnjo and Sellers, 1989). Serotonin-containing neurons are located predominantly in the raphe nucleus, and those neurons send projections throughout the CNS. Serotonin has been shown to modulate the activity of neurons in many regions of the CNS (cf. Weight and Salmoiraghi, 1968). The effect of ethanol on the firing of raphe neurons has been studied in vzzu~in unanesthetized, curarized rats by Chu (1984). In the dorsal raphe, two patterns of response were observed to the intraperitoneal administration of' 0.5-3 g/kg ethanol. In neurons that had a slow, regular firing pattern (0.5-5 Hz), ethanol produced a dose-dependent decrease in firing rate in 75% of the neurons. On the other hand, in neurons that had a more rapid firing rate (10-28 Hz), ethanol had no clear pattern of inhibition o r excitation. In the median raphe, the neurons exhibited an irregular pattern of discharge at frequencies that ranged from 2.5 to 35 Hz. T h e intraperitoneal administration of ethanol

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to median raphe neurons resulted in a reduction of spontaneous discharge frequency in 70%of the neurons. The effect of ethanol on neuronal firing in dorsal raphe nucleus has also been investigated by Verbanck et al. (1990). They found, in chloral hydrate-anesthetized rats, that with the intravenous infusion of 2 glkg ethanol, the rate of firing of approximately 50% of the dorsal raphe neurons was inhibited. In addition, the other 50% exhibited some excitation, but the excitation was preceded and followed by an inhibition of firing in most cells. The effect of ethanol on dorsal raphe neurons was studied in vitro in brain slices by Chu and Keenan (1987). They found that for cells with a slow discharge frequency (<5.9 spikeslsec), 22 mM ethanol increased the mean firing rate in 74%of the cells. With 43 mM ethanol, 60%of the neurons exhibited inhibition and 40% were excited. Higher concentrations of ethanol further increased the percentage of neurons showing inhibition, with 109, 217, and 434 mM ethanol inhibiting 75, 80, and 90%of the neurons, respectively. Ethanol in concentrations of 22 and 43 mM had little apparent effect on neurons with a fast discharge frequency (>6 spikedsec). Verbanck et al. (1990) have also studied the effect of ethanol on dorsal raphe neurons in brain slices. In their study, dorsal raphe neurons did not fire spontaneously, so that the experiments were conducted with a continuous superfusion of the a,-adrenergic agonist, phenylephrine (10 pM), to maintain a steady discharge. Under these conditions, ethanol concentrations between 10 and 50 mM produced concentration-dependent excitation of dorsal raphe neurons. To test whether the presence of phenylephrine was necessary to elicit this excitatory effect, they recorded from a neuron and confirmed the excitatory effect of 100 mM ethanol in the presence of phenylephrine; then both phenylephrine and ethanol were stopped. After the neuron had stopped firing, the slice was superfused with 100 mM ethanol alone. Without the presence of phenylephrine, ethanol produced only a transient excitation and further administration of ethanol was ineffective in eliciting additional excitation. Superfusion with phenylephrine restored the steady firing of the neuron and the sensitivity to ethanol.

F. OTHERCNS REGIONS In addition to the preceding studies wherein several different labs have investigated the effects of ethanol on a given brain region, there have also been several individual studies of ethanol effects on neurons in other brain regions, including the hypothalamus, spinal cord, septa1 area, and cerebral cortex.

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Because the lateral hypothalamus plays a role in the regulation of ingestive behavior and certain taste functions, Wayner et al. (1971) studied the effect of intravenously administered ethanol on the activity of lateral hypothalamic neurons. In urethane-anesthetized rats, 85% of the cells were reported to be sensitive to ethanol, with 53% excited by ethanol, 32% inhibited, and 15% not affected. In the spinal cord, Eidelberg and Wooley (1970) found that the intravenous administration of 0.1-0.6 g/kg ethanol decreased the spontaneous discharge in 96% of the dorsal horn interneurons studied in unanesthetized, spinalized or decerebrated cats. Only 1 of 24 interneurons studied increased its discharge rate following the administration of ethanol. On the other hand, Meyer-Lohmann et al. (1972) reported that the intravenous administration of 0.16-2.4 glkg ethanol caused a marked increase in the discharge of Renshaw cells. This effect was observed in anesthetized (pentobarbital or urethane-chloralose) as well as in unanesthetized, decerebrated o r spinalized cats. In a study of fetal mouse spinal cord neurons in tissue culture, Gruol (1982) found that the predominant effect of 10- 100 mM ethanol was a decrease in the spontaneous activity of the cells. Because Breese et al. (1988) had suggested that the medial septal area may be involved in ethanol-induced sedation, Givens and Breese (1990a) investigated the effect of ethanol on the firing rate of neurons in the septal area. In this study, the intraperitoneal administration of 0.75-3.0 g/kg ethanol suppressed neural firing of medial septal neurons in urethane-anesthetized and unanesthetized rats, but did not affect the neural activity of cells in the lateral septum. Collins and Roppolo (1 980) studied the effect of intravenously administered ethanol on neuronal activity in the primary somatosensory cortex of awake, nonparalyzed rhesus monkeys. They found that although pentobarbital ( 1-20 mg/kg) caused a profound, dose-dependent decrease in spontaneous activity, ethanol (0.1-2.5 g/kg) had little or no effect on the activity of somatosensory neurons.

G. COMMENTARY

'The preceding studies show that the administration of ethanol can affect the spontaneous firing rate of neurons in the central nervous system. These investigations indicate that in terms of spontaneous spike firing, ethanol does not appear to have a common action on all neurons. Indeed, a variety of different effects of ethanol have been observed. Comparison of the different studies suggests that a number of factors

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may contribute to diversity of neuronal responses to ethanol; these include the type of neuron studied, the preparation used, the method of ethanol administration, the rate and concentration of ethanol administered, and the presence or absence of anesthesia. Genetic factors also appear to be an important influence on the sensitivity of neurons to ethanol. In view of this diversity of influences, it appears that in order to attempt to elucidate the cellular and molecular basis of ethanol's action in the nervous system, it may be necessary to limit the number of variables that may affect neuronal responses to ethanol and to focus on ethanol's interaction with cellular and molecular mechanisms that are involved in the regulation of nerve cell activity. We will now turn our attention to studies on such mechanisms.

IV. Alcohol Effects on Cellular Mechanisms

At the cellular level, the nervous system can be considered to consist of two functional entities: (1) intrinsic electrical excitability of nerve cells, which is responsible for the generation of action potentials and patterns of action potential discharge, and (2) transmission at synapses, which involves both the release of the chemical neurotransmitter substance from presynaptic nerve terminals and the postsynaptic response to the action of neurotransmitter. These categories will serve as the organization framework for considering the cellular mechanisms of alcohol's actions in the nervous system.

A. ELECTRICAL EXCITABILITY AND VOLTAGE-GATED IONCHANNELS 1. Action Potential Generation and Membrane Properties Ethanol might affect the firing rate of neurons by altering the intrinsic electrical excitability of the membrane. A number of studies have examined the effects of ethanol on membrane excitability mechanisms. Early studies investigated ethanol effects on action potentials in the axons of squid (Armstrong and Binstock, 1964; Moore et al., 1964), lobster (Houck, 1969), and frog (Mullins and Gaffey, 1954; Quevedo et al., 1976). In general, they found that ethanol can reduce the rate of rise of the action potential, decrease action potential amplitude, and increase threshold, without appreciable change in membrane potential (<5 mV). However, in those experiments, the ethanol concentrations used were greater than 300 mM, concentrations that would be fatal in mammals.

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More recently, ethanol effects on action potential generation have been studied in intracellular experiments on mammalian neurons. Oakes and Pozos (1982a) investigated neurons from fetal rat dorsal root ganglia in tissue culture; ethanol was added to the culture medium for 1 hr and action potential parameters for ethanol-treated neurons were compared with those parameters for control neurons. They found that neurons treated with 11-65 mM ethanol did not have any significant difference in average resting membrane potential, rate of rise of the action potential, o r action potential amplitude; however, in cells treated with 33 and 65 IMethanol, the action potential had a slower rate of fall and an increased width. T h e authors suggested that a K + conductance was decreased as a result of a decrease in Ca2+ conductance. Gruol ( 1982) used intracellular recording techniques to study the effect of ethanol on fetal spinal cord neurons in tissue culture. She found that 10-100 mM ethanol had little or no effect on the amplitude of the action potential o r on membrane potential or membrane input resistance. T h e predominant effect observed was a decrease in the frequency of excitatory postsynaptic potentials (EPSPs) and inhibitory postsynaptic potentials (IPSPs), suggesting that ethanol exerts its action mainly through an interaction with synaptic processes. In a study of CA 1 pyramidal neurons in hippocampal slices using an interface chamber, Carlen et al. (1982) did not comment on the effect of ethanol on action potential amplitude, but reported that 5-20 mM ethanol, applied by drop application, increased the amplitude and duration of the afterhyperpolarization (AHP) that follows the generation of action potentials. i t was also reported that drop application of ethanol to slices on the surface of the bathing solution elicited a membrane hyperpolarization averaging 2.7 mV that was associated with a decrease in membrane input resistance. However, most of these observations have not been confirmed in subsequent studies using submerged slices and bath application of ethanol. Siggins et al. (1987) used submerged slices and bath application of ethanol to study the effect of ethanol on pyramidal neurons in the hippocampus. In CA1 pyramidal neurons, they found that spike amplitude was usually not affected by ethanol concentrations in the range of 10150 n a . In contrast to the observation by Carlen et al. (1982), ethanol usually had no effect o n the AHP (56%). In addition, ethanol decreased the amplitude of the AHP in 22% of the cells and slightly increased AHP amplitude in 22%. I n CA3 pyramidal neurons, 50 mM ethanol either produced no change o r reduced the magnitude of spontaneous AHPs by 10-24%, an effect that they felt caused an increase in the spontaneous firing rate. In this study, no consistent change in membrane potential or

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membrane input resistance in response to ethanol was observed. Benson et al. (1989) also studied ethanol effects on CA 1 pyramidal neurons using intracellular techniques. They did not report whether ethanol affects spike height, but they found that 100 mM ethanol does not affect the AHP. Because the AHP response was not altered by ethanol, they concluded that the inhibition of CA1 firing by ethanol is not mediated by an enhancement of the AHP. Niesen et al. (1988) used submerged hippocampal slices and bath application of ethanol to study dentate granule neurons from young (68 months) o r old (25-29 months) rats. They did not find any effect of 20 mM ethanol on membrane input resistance, action potential amplitude, or spike threshold. The spike AHP amplitude was not affected by ethanol in cells from young rats and was decreased in amplitude (1 mV) in cells from old rats. On the other hand, AHP duration was increased (0.5 sec) by ethanol in neurons from young animals, but AHP duration was not affected by ethanol in neurons from old animals. Membrane potential increased (3.3 mV) in association with ethanol treatment in neurons from young animals, but ethanol did not affect membrane potential in neurons from old animals. I n a number of recent voltage-clamp studies, ethanol, in concentrations of 100 mM or less, has not been found to activate a membrane current o r to cause a change of membrane conductance. The neurons studied include hippocampal CA3 pyramidal neurons in slices (Zbicz and Weight, 1982), chick spinal cord neurons in culture (Celentano et al. 1988), hippocampal neurons in culture (Lovinger et al., 1989), cortical neurons in culture (Aguayo, 1990), and adult dorsal root ganglion (DRG) neurons ((White et al., 1990).

2. Voltage-Gated Ion Channels Investigation of neuronal membrane physiology has revealed that voltage-gated ion channels underlie the generation of action potentials and the intrinsic electrical excitability of nerve cells (cf. Hille, 1984; Llinas, 1988). For technical reasons, earlier voltage-clamp experiments on the ion currents flowing through voltage-gated ion channels were conducted on giant invertebrate axons and neurons. More recently, the development of the single-electrode voltage-clamp and the patch-clamp techniques has permitted such studies in mammalian neurons. We will now turn our attention to studies on the effects of ethanol on voltagegated ion channels. a. Sodium Channels. Investigation of the effects of ethanol on the membrane conductance changes underlying action potential generation was undertaken initially in the squid giant axon. Moore et al. (1964) used

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the sucrose-gap technique to study the squid giant axon and found that 3% ethanol reduced the maximum conductance for sodium (gNa) by 1876, and 6% ethanol caused a 41% decrease of g;,. Ethanol had no significant effect on steady-state inactivation of gNdor the kinetics of gNa activation. Armstrong and Binstock ( 1964) reported similar observations using intraaxonal electrodes. As noted above, blood ethanol concentrations of this magnitude would be lethal in mammals. T h e effect of ethanol on the ion currents involved in action potential generation in Aplysia neurons has been studied by Bergmann et al. (1974). In this study, transient inward current was reduced in amplitude about 50% by 2% ethanol and 75%by 4%ethanol. Because the transient inward current in these neurons consists of both Na+ and Ca2+ currents, to isolate Na+ current, inward current was studied in a Ca2+-free external solution; under this condition, 4%ethanol caused a 50% reduction of Na+ current amplitude. In similar experiments on Aplysia neurons, Carnacho-Nasi and Treistman (1986) found that 300 mM ethanol reduced Na+ current by 37% and 500 mM ethanol decreased Na+ current by 58%. Mammalian sensory neurons acutely dissociated from their ganglia are particularly well suited for voltage-clamp analysis of ion currents in an adult mammalian neuron because of their spherical shape and absence of processes. We have studied and characterized Na+ currents in neurons acutely dissociated from rat nodose ganglion ((Ikeda et al., 1986) and in neurosecretory cell lines (PC12 and AtT20). In these cells, ethanol in concentrations of 100 mM o r less had no apparent effect on Na+ current (G. G. Schofield and F. F. Weight, unpublished observations). These observations are consistent with reports cited above that ethanol in concentrations of 100 mM or less does not significantly affect the amplitude o r rate of rise of action potentials in mammalian neurons. b. Potassium Channels. Potassium channels are known to play an important role in the repolarization of action potentials and the regulation of membrane excitability. Several types of potassium currents have been characterized (cf. Rudy, 1988) and will be considered separately. i. Delayed rectifier current. T h e slow outward K + current that is responsible for action potential repolarization in invertebrate and vertebrate axons, as well as many neurons, has been called the delayed rectifier current. In voltage-clamp experiments on squid axon using an intraaxonal electrode, Moore (1966) reported that 3.2%ethanol had no significant effect on delayed rectifier K + current. Using a similar technique to study squid axon, Armstrong and Binstock (1964) reported similar observations. On the other hand, using the sucrose-gap recording technique to study the squid axon, Moore et al. (1964) found that 3%

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ethanol reduced the maximal membrane conductance for potassium (gk)by 20%, whereas 6% ethanol decreased gK by 31%. In two-electrode voltage-clamp experiments on Aplysia neurons, Bergmann et al. (1974) observed that although 4% ethanol reduced Na+ current amplitude by 50%, it did not significantly affect the amplitude of delayed rectifier K + current. T h e neurosecretory cells of the pineal gland are particularly well suited for studying K + currents in mammalian cells, because of the absence of other currents. We have investigated the K + currents present in cells acutely dissociated from adult rat pineal gland using the whole cell patch-clamp technique and characterized a delayed rectifier current similar to that descried in squid axon (Aguayo and Weight, 1988). In these cells, ethanol had no significant effect on delayed rectifier K + current in concentrations of 100 mM or less (L. G. Aguayo and F. F. Weight, unpublished observations). ii. A-current. A low-threshold, transient outward K current found in many invertebrate and vertebrate neurons has been called A-current. This current appears to be important in reducing the frequency of repetitive spike firing. In a study of A-current in Aplysia neurons, Bergmann et al. (1974) found that 4% ethanol produced a 20% reduction in the amplitude of A-current in the cells they studied that exhibited this current ( L 2 to L-6 and R-15). Camacho-Nasi and Treistman (1986) also studied the effect of ethanol on A-current in Aplysia neurons and found that ethanol (200-600 mM) slowed the A-current decay time course in the giant metacerebral cell (MCC). In a subsequent study, Treistman and Wilson (1987a) showed that 200-600 mM ethanol increased the A-current decay time constant in two cell types (MCC and R15), but did not significantly affect the decay in a third cell type (Bl). Because the longer chain-length alcohols, butanol and hexanol, did not have similar effects on A-current decay kinetics (Treistman and Wilson, 1987b) and there was no significant difference in the temperature sensitivity of A-current decay in ethanol-sensitive and ethanol-insensitive neurons (Treistman and Grant, 1990), the authors suggested that it is unlikely that ethanol exerts its action on A-current by perturbation of the bulk phase of membrane lipid. We have characterized A-current in neurosecretory cells acutely dissociated from the pineal gland of adult rats using the whole cell patchclamp recording technique (Aguayo and Weight, 1988). In these cells, ethanol did not significantly affect A-current amplitude or decay kinetics in concentrations of 100 mM or less (L. G. Aguayo and F. F. Weight, unpublished observations). iii. M-current. M-current is a slowly activating, sustained outward +

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K' current that is inhibited by acetylcholine acting on muscarinic receptors (hence the name), as well as by several other neurotransmitters. Inhibition of M-current results in an increased membrane excitability. Moore et al. (1990) investigated the effect of ethanol on M-current in CA 1 pyramidal neurons in hippocampal slices using the single-electrode voltage-clamp technique. They found that 22 mM ethanol reduced the amplitude of M-current by 2270, and 44 mM ethanol produced a 36% reduction of M-current amplitude. T h e effect was reversible in some but not all cells. T h e reduction of M-current was not blocked by atropine, indicating that it is not due to a presynaptic release of acetylcholine acting on postsynaptic muscarinic receptors. Moore et al. (1990) suggested that the effect of ethanol on M-current may contribute to the interaction of ethanol with acetylcholine and somatostatin effects on spike firing. In extracellular experiments recording from hippocampal pyramidal neurons, ethanol had been found to enhance the excitatory response to acetylcholine and the inhibitory response to somatostatin (Mancillas et ul., 1986), and in single-electrode voltage-clamp experiments, acetylcholine inhibits M-current (Brown, 1988) and somatostatin augments M-current (Moore et al., 1988). On the other hand, in whole cell patch-clamp experiments on bullfrog sympathetic neurons, C. Li and F. F. Weight (unpublished observations) and G. White and F. F. Weight (unpublished observations) were unable to detect any effect of ethanol (100 m M or less) on M-current. iv. Inward rect$el- current. Some neurons show a pronounced increase in inward current at membrane potentials negative to the K + equilibrium potential ( E K ) .This current has been called the inward (or anomalous) rectifier; it results from an increase in K + current, which is inward at potentials negative to E,. Shefner and Osmanovic (1988) studied the effect of ethanol on inward rectifier current in single-electrode voltage-clamp experiments on locus coeruleus neurons in brain slices. They found that 30-200 mM ethanol shifted, in the depolarizing direction, the potential range over which inward rectification occurred. This effect was similar to the shift of voltage dependence of inward rectification observed when extracellular K + concentration was increased. T h e shift observed with 200 mM ethanol was equivalent to the shift resulting from an increase of extracellular K + concentration of 0.9 mM. The authors suggest that this shift results from an elevation of K + concentration in the extracellular space immediately outside the neuronal membrane. c. Calcium Channels. Calcium channels play an important role in the regulation of nerve cell excitability and the release of neurotransmitters (cf. Augustine et ul. 1987; Tsien, 1987). In invertebrate neurons,

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Bergmann et al. (1974) studied the effect of ethanol on the Ca2 component of the action potential in Aplysza neurons; the Na+ component of the action potential was inhibited by tetrodotoxin (TTX) or Na+ -free external solution. They found that 4% ethanol inhibited the Ca2+ current by about SO%, an inhibition similar to that of Na+ current in the same neurons. Schwartz (1985) also studied the effect of ethanol on Ca2+ current in Aplysia neurons (L2-L4, L6, and R15) and found that 4% ethanol reduced Ca2+ current amplitude by about 50%. This effect was not blocked by intracellular injection of EGTA. Ca2+ current appeared to be more sensitive to the effects of ethanol in a study of Aplysia neurons (R15, R2, L2, and LPl) by Camacho-Nasi and Treistman (1986). In their study, 50, 200, and 400 mM ethanol inhibited Ca2+ current by 19, 55, and 76%, respectively. A more detailed analysis revealed that the ethanol inhibition of Ca2+ current developed over a period of 20 min; in some cases the inhibition was only partially reversible, and ethanol appeared to augment Ca2+ -dependent inhibition of Ca2 current (see below) because the effect was not observed with Ba2 substitution for extracellular Ca2 (Camacho-Nasi and Treistman, 1987). T h e effect of ethanol on Ca2+ current was also investigated in Helix neurons by Oyama et al. (1986). In these neurons, ethanol produced a concentration-dependent reduction of Ca2 current amplitude over a concentration range from 178 to 730 mM; a concentration of 178 mM ethanol reduced Ca*+ current by about 15%, and 730 mM reduced Ca2+ current by about 50%. This concentration range of ethanol also increased the decay rate of Ca2+ current. Ethanol reduced the peak amplitude of Ca2 current without shifting the current-voltage relationship and with little effect on steady-state inactivation. In mammalian neurons, Oakes and Pozos (1982b) used intracellular recording to study fetal rat dorsal root ganglion neurons in tissue culture. The external solution contained high Ca2+ (5.4 mM) and 3.0 mM Ba2+, to inhibit K + conductances and increase the width of the action potential. Under these conditions, 11-109 mM ethanol produced a concentration-dependent reduction in spike width. The authors interpreted this effect as due to a reduction in Ca2+ conductance. I n the same type of preparation, Eskuri and Pozos (1987) observed that although a moderate change of temperature (27-43°C) affected the potency of ethanol in reducing spike width, moderate temperature change alone did not significantly affect spike width. Because temperature change in this range has been shown to alter membrane fluidity, the authors concluded that the effect of ethanol on spike width is not due to a nonspecific disturbance of membrane fluidity. Llinas (1988) and Scott et al. (1990) have reported that 1-octanol (20 +

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and 1 pV, respectively) can selectively inhibit low-threshold, transient (T-type) Ca2+ current in inferior olive and cultured DRG neurons, respectively. On the other hand, Twombly el al. (1990) studied ethanol effects on two types of voltage-activated (;a2+ currents in N1E-115 neuroblastoma and NG108- 15 neuroblastoma x glioma hybrid cells. Ethanol concentrations of 100 and 300 mM reduced Ca2 current amplitude by approximately 15 and 40%, respectively. Concentration-response curves were similar for both low-threshold, transient (T-type) and high-threshold, long-lasting (L-type) Ca2 currents in both NG108- 15 and N 1E-115 cells. Ethanol produced a mild acceleration in the decay kinetics of both current types; however, the ethanol-induced reduction of Ca2 current amplitude was not associated with an alteration in the voltage dependence of activation or inactivation. In addition, the current reduction did not appear to be use dependent. T h e effect of ethanol on Ca2+ currents in hippocampal dentate granule neurons has been investigated by Carlen at al. (1991) using the single-electrode voltage clamp. At room temperature (20-23"C), 20 mM ethanol reduced the amplitude of high-threshold, inactivating Ca2 current (N-type) by 33% and high-threshold, long-lasting Ca2+ current (Ltype) by 55%. Similar effects were observed with 50 mM ethanol. A reduction in Ca2 current amplitude by ethanol was not observed when the Cay chelator EGTA was injected intracellularly or when recording at 30°C. T h e authors propose that the effects of ethanol are associated with an elevation of intracellular Ca2 that secondarily inhibits voltageactivated Ca2 current. We have used the whole cell patch-clamp technique to characterize Ca2+ currents and study their regulation in AtT20 cells (Lewis et al., 1986; Lewis and Weight, 1988), and in neurons freshly dissociated from adult rat nodose ganglion (Ikeda et al., 1986) and adult rat dorsal root ganglion (White el al., 1989). Using solutions in the patch pipette (which dialyze the interior of the cell) containing 10 mM EGTA, we did not observe effects of ethanol, in concentrations of 100 mM or less, on lowthreshold, transient (T-type) o r high-threshold, long-lasting ( h y p e ) Ca2+ currents in the nodose neurons (S. R. Ikeda and F. F. Weight, unpublished observations), the dorsal root ganglion neurons (G. White and F. F. Weight, unpublished observations), or the AtT20 cells (D. L. Lewis and F. F. Weight, unpublished observations). Ethanol has been shown to cause a release of Ca2 from or an elevation of intracellular Ca2+ in brain synaptosomes (Pande et al., 1983; Daniel1 el al., 1987; Davidson et al., 1988; Rezazadeh et al., 1989) and clonal neural cells (Rabe and Weight, 1988). This effect appears to be due to a release of Ca2+ from intracellular stores because (1) it can be elicited in a Ca2+-free extracellular medium (Pande et al., 1983; Re+

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ALCOHOL ACTIONS IN T H E NERVOUS SYSTEM

zazadeh et al., 1989); ( 2 ) it can be blocked by dantrolene (Pdnde et al., 1983), a drug that blocks Ca2+ release from endoplasmic reticulum of neurons (Kuba, 1980; Weight and McCort, 1981); and (3) ethanol can release Ca2+ from isolated brain microsomes (Shah and Pant, 1988; Daniel1 and Harris, 1989; Machu et al., 1989).Because elevation of intracellular Ca2+ has been found to inhibit Ca2+ current in a number of excitable cells (Brehm and Eckert, 1978; Tillotson, 1979; Standen, 1981; Pitler and Landfield, 1987) and there was little or no buffering of intracellular Ca2 in those experiments where pharmacological concentrations of ethanol have been reported to inhibit Ca2+ current, it is possible that the reduction in Ca2+ current by pharmacological concentrations of ethanol may be secondary to an elevation of intracellular Ca2+. d . Commentary. The preceding discussion indicates that ethanol can reduce the action potential and voltage-gated Na+ and K + currents in both invertebrate and vertebrate neurons. However, in most cases, the ethanol concentrations that produce significant inhibition of Na and K + currents are concentrations that would be fatal in mammals. Voltage-gated Ca2 current can also be reduced in amplitude by ethanol in both invertebrate and vertebrate neurons. Although in a number of studies the ethanol concentrations that significantly inhibit Ca2 current would be lethal for mammals, there have been a few reports of Ca2+ current inhibition by ethanol concentrations in the pharmacological range. Because ethanol can elevate intracellular Ca2+ and intracellular Ca2+ elevation can inhibit Ca2+ current, further work is needed to determine whether the reduction of Ca2 current resulting from pharmacologically relevant concentrations of ethanol is secondary to an elevation of intracellular Ca2+. In addition, selective effects of l-octanol have been reported on T-type Ca2+ current. Because, in some cases, long-chain alcohols have been found to have effects on ion channels that differ from those of short-chain alcohols (e.g., Pennefather and Quastel, 1980; Bradley et al., 1984; Treistman and Wilson, 1987b), further work is needed to determine whether the effect of l-octanol is a unique property of long-chain alcohols or there are types of Ca2 channels in some neurons that are particularly sensitive to ethanol. +

+

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B. SYNAPTIC TRANSMISSION AND TRANSMITTERGATED IONCHANNELS 1. Excitatory Transmission and Glutamate-Gated Ion Channels There have been relatively few studies of ethanol effects on excitatory synaptic potentials in the CNS. In the spinal cord of pentobarbitalanesthetized cats, Klee et al. (1975) found that ethanol in i.v. doses of 1.5

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g/kg reduced the amplitude of EPSPs recorded in motoneurons for both mono- and polysynaptic pathways, with polysynaptic pathways being more sensitive. In hippocampus, Doller et al. (1980), using submerged slices, found that 60-300 n M ethanol produced a concentration-dependent depression of synaptically evoked responses in CA 1 pyramidal neurons and dentate granule cells. On the other hand, Carlen et al. (1982), using slices on the surface of the bathing fluid (interface chamber), reported that 10 or 20 mM ethanol augmented EPSP amplitude in CAI pyramidal neurons. However, this observation has not been confirmed by subsequent studies using submerged slices. In a more recent study from the same laboratory, Niesen et al. (1988) did not find any effect of 20 mM ethanol on EPSP amplitude or half-width in dentate granule neurons. Siggins et al. (1987) observed that ethanol in concentrations from 10 to 350 mM most often reduced the amplitude of EPSPs in both CA1 and CA3 pyramidal neurons. Recently, Lovinger et al. (1990b) found no significant effect of 50 mM ethanol on population EPSPs (pEPSPs) in CA1 pyramidal neurons, but 100 mM ethanol produced a 9% reduction in pEPSP amplitude that was statistically significant (see below). In recent years, L-glutamate has come to be recognized as the principal mediator of fast excitatory neurotransmission in the mammalian CNS (cf. Mayer and Westbrook, 1987; Collingridge and Lester, 1989). Studies on the postsynaptic action of glutamate have revealed that it activates at least three types of ligand-gated ion channels, designated by the agonists that activate them-N-methyl-D-aspartate (NMDA), kainate, and quisqualate [or a-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA)] (cf. Mayer and Westbrook, 1987; Collingridge and Lester, 1989). Recent studies indicate that ethanol has significant effects on these ion channels. T h e effects of ethanol on glutamate-gated ion channels will be reviewed below. a. NMDA Channels. Excitatory postsynaptic potentials mediated by NMDA receptors have a slower time course than do the fast EPSPs mediated by the non-NMDA excitatory amino acid receptors kainate and quisqualate (cf. MacDermott and Dale, 1987). Although the nonNMDA EPSPs can be recorded at negative membrane potentials, some depolarization of the postsynaptic neuron may be needed to observe NMDA receptor-mediated EPSPs, because at negative membrane potentials Mg2+ blocks the NMDA-gated ion channel and this block is relieved by depolarization (Nowak et al., 1984; Mayer et al., 1984). The NMDA channel also differs from the non-NMDA channels in that the NMDA channel is permeant to Ca2+ (MacDerinott et al., 1986) and the presence of glycine appears to be essential for gating of the channel by agonists (Johnsonand Ascher, 1987; Kleckner and Dingledine, 1988). There has

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been considerable recent research interest in NMDA channels, because it has been suggested that they may be involved in CNS processes such as long-term potentiation (Gustafsson and Wigstrom, 1988), certain types of learning (Morris et al., 1984), neural development (Cline et al., 1987; Kleinschmidt et al., 1987; Rauschecker and Hahn, 1987), neurotoxicity (Rothman and Olney, 1987), and kindling and epilepsy (Dingledine et al., 1986; Pate1 et al., 1988). There have been several recent studies of ethanol effects on NMDA channels. In whole cell patch-clamp experiments on adult rat dorsal root ganglion neurons, Lovinger and Weight (1988) and White et al. (1990) found that ethanol in concentrations from 2.5 to 50 mM produced a concentration-dependent inhibition of NMDA-activated ion current (Fig. 1). T h e ethanol concentration that produced 50% inhibition (IC5,) was 10 mM, and maximal inhibition of NMDA-activated current (83%) was observed with an ethanol concentration of 50 mM (Fig. 1A and C). The inhibition produced by 100 mM ethanol was not significantly different than the inhibition by 50 m M ethanol (Fig. 1C). In addition, the NMDA-activated response was inhibited throughout the duration of ethanol application with ethanol applications up to 5 min in duration (Fig. 1B). I n some neurons, ethanol concentrations of 1 mh4 and 500 pkf were observed to potentiate NMDA-activated current; ethanol concentrations of 250, 100, and 50 pJ4 had no apparent effect on NMDA-activated current. Because these neurons did not have kainate- or quisqualateactivated currents, the effect of ethanol was also studied in hippocampal neurons, which are known to have all three types of glutamate-activated ion channels. T h e effect of ethanol on glutamate-activated ion channels in cultured mouse hippocampal neurons was studied by Lovinger et al. (1989) using the whole cell patch-clamp technique. As in DRG neurons, ethanol also greatly reduced the amplitude of NMDA-activated current in hippocampal neurons (Fig. 2A, left). The inhibition was concentration dependent over the range 5-50 mM ethanol (Fig. 2A, right). The IC,, for inhibition of NMDA-activated current was 30 mM. The average inhibition by 50 mM ethanol was 61%, and the inhibition by 100 mM ethanol was not significantly different than that by 50 mM. An inhibition of NMDAactivated current was not observed with 2.5 mM ethanol in hippocampal neurons; in some cells, however, this concentration potentiated NMDAactivated current. Application of 100 mM ethanol alone did not activate any membrane ion current or alter membrane input resistance, indicating that the ethanol-induced reduction of NMDA-activated current was not due to nonspecific membrane actions or alteration of whole cell recording conditions. Inhibition of NMDA responses by ethanol has also been observed in

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other brain regions and experimental conditions. Lovinger et al. (1990a) found that ethanol produced a similar inhibition of NMDA-activated current in cultured neurons from hippocampus, spinal cord, and neocortex. In addition, the inhibition was similar in neurons internally dialyzed with solutions containing CsC1, KCl, or KMeSO,. Using outsideout tear-off patches from cultured hippocampal neurons, LimaLandman and Albuquerque (1989) reported that ethanol concentrations from 1.74 to 8.65 mM increased the probability of channel opening without changing mean channel open time, whereas ethanol concentrations from 87 to 174 mM decreased both the probability of channel opening and the mean channel open time. Recording extracellularly from neurons in the medial septa1 nucleus, Simson et al. (199 1) recently reported that the intraperitoneal administration of 0.75 to 2.5 g/kg ethanol can inhibit NMDA-activated spike firing in vivo. In biochemical experiments, ethanol has been found to reduce NMDA-induced Ca2 influx and cyclic GMP production in cultured cerebellar granule cells (Hoffman et al., 1989), neurotransmitter release from slices of neocortex (Gothert and Fink, 1989; Gonzales and Woodward, 1990) and striatum (Woodward and Gonzales, 1990), and intracellular Ca2 elevation in dissociated brain cells (Dildy and Leslie, 1989). T h e concentration range over which increasing concentrations of ethanol produced increasing inhibition of NMDA-activated current in hippocampal neurons (5-50 mM) corresponds to the clinical bloodethanol concentrations associated with increasing impairment of motor and cognitive function (viz. intoxication) (Kissin, 1988), raising the question of whether the inhibition of NMDA-activated current by ethanol may be responsible for the behavioral manifestations of intoxication. Because different alcohols differ in their potency for producing intoxication, Lovinger et al. (1989) studied the potency of different alcohols for inhibiting NMDA-activated current. Figure 3 shows that 200 mM methanol (Fig. 3A, left), 10 mM 1-butanol (Fig. 3B, left), and 0.5 mM isopentanol (Fig. 3C, left) elicited an inhibition of NMDA-activated cur+

+

FIG. l. Ethanol inhibition of N-methyh-aspartate (NMDA)-activatedion current. (A) Effect of 50 mM ethanol (EtOH) on current elicited by application of NMDA. (B) Graph plotting amplitude of current activated by NMDA as a function of time, and illustrating two periods of exposure to ethanol by bath application. (C) Average percent inhibition of NMDA-activated current as a function of ethanol concentration. Each point represents mean SD percent inhibition observed in at least four neurons. Whole cell patch-clamp recordings from neurons freshly dissociated from adult rat dorsal root ganglia. In A, B, and C, 100 pM NMDA was applied from a macropipette by diffusion. In A and C, ethanol was also in the macropipette, as indicated. The bar above each record denotes time of drug application. From White et al. (1990).

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FIG.2. El hano1 inhibition of excitatory amino acid-activated ion currents in hippocanipal neurons. (A) (Left) Effect of 50 rmlJ ethanol on current elicited by application of50 phi’ NMI)A. (Right) Average percent inhibition of NMDA-activated ion current as a function of ethanol concentration. (B) (Left) Effect of 50 nuCf ethanol on current elicited by application of 1 0 k%Jkainate. (Right) Average percent inhibition of kainate-activated ion current as a turiciion of eLhanol concentration. (C) (Left) Effect of 50 ml.1 ethanol on current elicited by application of 1 ~ t . 1quisqualate. (Right) Average percent inhibition of quisqualateactivated ion current as a f-unction of ethanol concentration. Whole cell patch-clamp recording from cultured mouse hippocampal neurons. Excitatory amino acids and ethanol

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ALCOHOL ACTIONS IN THE NERVOUS SYSTEM

rent that was comparable to the inhibition by 50 mM ethanol (Fig. 2A, left). Concentration-response curves revealed that the alcohols differ in their potency for inhibiting NMDA-activated current over a range of concentrations. Methanol was the least potent alcohol tested, inhibiting NMDA-activated current with a threshold of 25 mM and an IC,, of 117 mM (Fig. 3A, right). On the other hand, 1-butanol was more potent than ethanol, with a threshold of 0.01 mM and an IC,, of 1.14 mM (Fig. 3B, right). The most potent alcohol tested was isopentanol, which had a threshold of 0.001 mM and an IC,, of 0.32 mM (Fig. 3C, right). Because different alcohols exhibit different degrees of hydrophobicity, Lovinger et al. (1989) examined the relationship between the potency of different alcohols for inhibiting NMDA-activated current and their hydrophobicity. Figure 4A illustrates that as the hydrophobicity of the alcohols increased, the IC,, for inhibition of NMDA-activated current decreased. In addition, there is a significant linear relation between these parameters, suggesting that the potency of different alcohols for inhibiting NMDA-activated current increases as a function of increasing hydrophobicity. Different alcohols also differ in their potency for producing intoxication, so that Lovinger et d.(1989) also determined the relationship between the potency of different alcohols for inhibiting NMDA-activated current and their potency for producing intoxication. Figure 4B shows that as the IC,, for inhibition of NMDA-activated current decreased, the ED3 for inloxication decreased. Moreover, there is a significant linear relationship between these parameters, suggesting that the more potent the alcohol is in inhibiting NMDA-activated current, the greater its potency for producing intoxication. The mechanism of the ethanol-induced inhibition of NMDAactivated ion current has been investigated by White et al. (1992). One mechanism that might account for the reduction of current amplitude is a voltage-dependent block of the channel similar to that produced by phencyclidine (PCP) or Mg2+. They found, however, that the percent inhibition of peak NMDA-activated current by 50 mM ethanol was similar over a range of membrane potentials from -60 to +60 mV. A second possible mechanism that could account for the current reduction is an alteration of the ion selectivity of the channel. When tested, ~

~

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~

were applied from a macropipette. In A, the extracellular medium contained no added Mg2+; in B and C, the extracellular medium contained 1 mM Mg*+ and 50 (uz.I D-2amino-5-phosphonovalerate. All records were taken at a membrane potential of -50 mV. The bar above each record indicates time of drug application. Each point in the graphs represents the means 2 SEM percent inhibition observed in at least four neurons. From Lovinger et al. ( 1 989).

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FIG.:1. Inhibition of NMDA-activated ion current hy different alcohols. (A) (Left) Effect of 200 nuEl methanol o n current elicited by 50 )uM NMDA. (Right) Average percent inhibition o f NMDA-activated current as a function of methanol concentration. (8)(Left) Effect o f 10 n&f 1 -butand on current elicited by 50 pM NMDA. (Right) Average percent inhibition of NMDA-activated cirrr-ent as a function of I-butanol concentration. (C) (Left) Elfect of 0.5 1niL1 isopent;rriol o n current elicited hy 50 ph4 NMDA. (Right) Average percent inhibition of NMDA-activated current as a function of isopentanol concentration. Whole cell patch-clamp recording from cultured mouse hippocampal neurons. Experimental conditions as in Fig. 2. From Lovinger r! al. (1989).

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however, 50 mM ethanol did not aIter the reversal potential of the NMDA-activated current. A third possibility is that ethanol might alter the affinity of the NMDA-binding site. However, the percent inhibition by 10-100 mM ethanol did not differ with NMDA concentrations from 10 to 100 cuz/r. Fourth, ethanol might interact with one of the binding sites on the NMDA receptor/ionophore complex that can regulate NMDAactivated current; these sites include the binding sites for glycine, Mg2 , Zn2+, and PCP/ketamine (cf. Wong and Kemp, 1991). They found that the percent inhibition by 50 mM ethanol did not differ with concentrations of glycine from 10 nM to 1 cuz/r (but see also Woodward and Gonzales, 1990; Rabe and Tabakoff, 1990; Peoples and Weight, 1991a), o r in the absence or presence of different concentrations of Mg2+ (10, 100, and 500 pM), Zn2+ (5 and 20 cuz/r), or ketamine (2 and 10 cuz/r). The observations indicate that ethanol does not reduce NMDA-activated current by a voltage-dependent block of the ion channel or by altering the ion selectivity of the channel. The data also provide evidence that ethanol does not appear to inhibit NMDA-activated current by interacting with several previously described binding sites on the NMDA receptor/ ionophore complex. b. Kainate and Quisqualute Channels. Kainate- and quisqualateactivated channels appear to mediate the majority of fast EPSPs in the CNS. T h e effect of ethanol on kainate- and quisqualate-activated currents has been studied by Lovinger et al. (1989) in cultured hippocampal neurons using the whole cell patch-clamp technique. In the same neurons that intoxicating concentrations of ethanol produced a marked inhibition of NMDA-activated current, they found that kainate- and quisqualate-activated currents were relatively insensitive to ethanol concentrations less than 50 mM. Figure 2 (left) illustrates a comparison of the effects of 50 mM ethanol on NMDA-, kainate-, and quisqualateactivated currents. As can be seen, although 50 mM ethanol markedly reduced the amplitude of NMDA-activated current (Fig. 2A, left), this concentration of ethanol had a relatively small effect on kainate- and quisqualate-activated current (Fig. 2B and C, left). On average, 50 mM ethanol elicited a 61% inhibition of NMDA-activated current; by contrast, the reduction of kainate- and quisqualate-activated current by this ethanol concentration was only 18 and 15%, respectively. Similarly, although 25 mM ethanol inhibited NMDA-activated current by more than 50%, the reduction of kainate- and quisqualate-activated current by 25 mM ethanol was less than 10%. The concentration-response curves for ethanol inhibition of these ion currents are shown in Fig.2 (right). Comparison of these curves reveals that although the inhibition of NMDAactivated current did not increase significantly when ethanol concentration was increased from 50 to 100 mM, the inhibition of kainate- and +

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ALCOHOL ACTIONS IN THE NERVOUS SYSTEM

32 1

quisqualate-activated currents continued to increase significantly when ethanol increased from 50 to 100 mM. In fact, the inhibition of nonNMDA glutamate-activated current continued to increase at ethanol concentrations greater than 100 mM, so that at a concentration of 200 mM ethanol the inhibition was greater than 45% (Fig. 5). c. NMDA Receptor-Mediated Synaptic Excitation. In order to determine whether ethanol would have an action on NMDA receptor-mediated synaptic excitation that was similar to its effect on NMDA-activated ion current, Lovinger et al. (1990b) studied the effect of ethanol on NMDA receptor-mediated pEPSPs in hippocampal slices from the adult rat. Figure 6 illustrates the effect of ethanol on NMDA receptor-mediated synaptic responses in CA 1 pyramidal neurons. The records in Fig. 6A show that 50 mM ethanol reversibly reduced the amplitude of the NMDA receptor-mediated pEPSP by nearly half. It can also be seen in these records that this concentration of ethanol did not significantly affect the amplitude of the presynaptic fiber volley. Figure 6B plots graphically the time course of the effect of 50 mM ethanol on NMDA receptor-mediated pEPSP amplitude. As can be seen, pEPSP amplitude began to decrease when ethanol reached the bath. Within minutes after the onset of ethanol application, the reduction in pEPSP amplitude stabilized at a maximal level. T h e pEPSP gradually returned to control amplitude over several minutes when ethanol was washed from the recording chamber. The average reduction of NMDA receptor-mediated pEPSP amplitude by 50 mM ethanol was 43%. A similar magnitude of NMDA receptormediated pEPSP inhibition was observed over the duration of ethanol applications lasting as long as 2.5 hr. Moreover, full recovery from the effects of ethanol was observed, even after such prolonged applications. Figure 6C illustrates that 50 mM ethanol greatly reduced the amplitude of population spikes activated by NMDA receptor-mediated EPSPs, as would be expected when the EPSPs that trigger action potential generation are reduced in amplitude.

FIG.4. Correlation between the potency of different alcohols for inhibiting NMDAactivated ion current, the hydrophobicity of the alcohols, and the potency of the alcohols for producing intoxication. (A) Log-log graph plotting IC5,, of four alcohols tested for inhibition of NMDA-activated current (Figs. 2 and 3) as a function of their membranebuffer partition coefficients. The linear relation between IC5,, for inhibition of NMDAactivated current and membrane-buffer partition coefficient has a slope of - 1.30 2 0.055 (p < 0.01). (B) Log-log graph plotting ED3 for intoxication by four alcohols versus the IC50 for inhibition of NMDA-activated by the alcohols (Figs. 2 and 3). The linear relation between the IC5,,for inhibition of NMDA-activated current and the ED3 for intoxication has a slope of 1.579 +- 0.099 (p < 0.025). From Lovinger et al. (1989).

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T h e bar graphs in Fig. 7 illustrate the average inhibition of NMDA receptor-mediated pEPSPs by several ethanol concentrations from 1 to 100 mM. The effect of 1 mM ethanol was not statistically significant. However, ethanol concentrations of 25 mM and above produced significant inhibition of NMDA receptor-mediated pEPSPs. At 25 mM, ethanol inhibited these synaptic potentials by 25%, and 50 mM ethanol inhibited these pEPSPs by 43%. The inhibition by 100 mM ethanol was not significantly different from that observed with 50 mM ethanol. Thus, the concentration range over which ethanol inhibited NMDA receptormediated pEPSPs was roughly similar to the concentration range over which ethanol inhibits NMDA-activated ion current in cultured hippocampal neurons (Lovinger et al., 1989) and isolated dorsal root ganglion neurons (White et al., 1990). In addition, 50 mM methanol inhibited NMDA receptor-mediated pEPSPs by 1796, which was significantly less than the inhibition by 50 mM ethanol, and 1 mM 1-butanol inhibited these synaptic responses by 27%, which was significantly greater than the effect of 1 mM ethanol. These observations correspond to the effect of different alcohols on NMDA-activated ion currents (Lovinger et al., 1989). d . Non-NMDA Receptor-Mediated Synaptic Excitation. To determine whether ethanol affects synaptic excitation mediated by non-NMDA glutamate receptors in a manner that corresponds to its effects on the currents activated by the glutamate agonists, kainate and quisqualate, Lovinger et al. (1990b) also studied the effect of ethanol on non-NMDA glutamate receptor-mediated pEPSPs in the hippocampal slice. Figure 8A illustrates that 50 mM ethanol had little or no effect on pEPSPs mediated by non-NMDA glutamate receptors. On average, the effect of 50 mM ethanol on non-NMDA glutamate receptor-mediated pEPSPs was not statistically significant (Fig. 8B). On the other hand, in the presence of 100 mM ethanol, non-NMDA glutamate receptor-mediated pEPSPs were reduced 9% in amplitude, which was a statistically significant reduction of pEPSP amplitude (Fig. 8B).

FIG. 5. Ethanol inhibition of non-NMDA excitatory amino acid-activatedion currents. (A) Graph plotting percent inhibition of non-NMDA receptor-mediated glutamate current as a function of ethanol concentration. Data are from cultured mouse hippocampal neurons. (B) Graph plotting percent inhibition of ion current activated by kainate as a function of ethanol concentration. Data are from neurons freshly dissociated from adult rat hippocampus. In both A and B, the values are mean ? SEM, the recording utilized the whole cell patch-clamp method, and the external solution contained 1 mM Mg2+ and 50 fl D-2amino-5-phosphonovalerate.From Lovinger et al. (1990a).

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FIG.ti. Ethanol inhibition of NMDA receptor-mediated synaptic potentials and synaptically activated action potentials in hippocampus. (A) Effect of 50 nlM ethanol on NMDA rereptor-mediated population excitatory postsynaptic potential (pEPSP). (B) Graph plotting amplitude of individual pEPSPs in A, as a function of time. Bar indicates time of 50 mM ethanol superfusion of hippocampal slice. (C) Effect of 50 mM ethanol on population spike activated by NMDA recei""r-riiediatcd EPSPs. Responses elicited in rat hippocampal slice by stimulation of Schaffer collateral/comniissural pathway in extracellular medium and 0.1 mhl M$+. Responses in A containing 10 pVf 6,7-dinitro-quinoxaline-2,3-ciione and B recorded in CAI stratum pyramidale, and responses in C; recorded in CAI stratum pyramidale. From Lovinger rt al. (1990b).

e. Cornmentar). Ethanol has been found to produce a concentrationdependent inhibition of NMDA current over the concentration range 550 mM. This corresponds to the blood-alcohol concentration range over which intoxication occurs in nontolerant humans (Kissin, 1988). Increasing sedation and signs of deeper stages of general anesthesia are

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50 6o

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FIG. 7. Concentration dependence of ethanol inhibition of NMDA receptor-mediated pEPSPs in hippocampus. Bar graphs show average percent inhibition (mean f SEM) of NMDA receptor-mediated pEPSPs by 1, 25, 50, and 100 mM ethanol; * , p < 0.025 versus baseline and 1 mM ethanol; **, p < 0.05 versus baseline, 1, and 25 mh4 ethanol. Recording from rat hippocampal slice. Experimental conditions as in Fig. 6. From Lovinger et al. (1990b).

observed with progressive increases in blood-ethanol concentration greater than 200-250 mg%. Because the inhibition of NMDA current by ethanol did not increase significantly at concentrations greater than 50 mA4, the general anesthetic effects of ethanol would not appear to be due to inhibition of responses to NMDA. On the other hand, ethanol concentrations greater than 50 mA4 produce progressive inhibition of kainate and quisqualate currents. Because kainate and quisqualate receptors mediate most fast EPSPs in the CNS, inhibition of these responses would be expected to produce general CNS depression.

2. Inhibitory Transmission and GABA-Gated Zon Channels There have been relatively few reports of ethanol effects on inhibitory synaptic transmission in the CNS, but there have been a number of studies on the effect of ethanol on responses to the major inhibitory transmitter in the brain, y-aminobutyric acid (GABA), with a number of

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different experimental results. In early studies, Miyahara et al. (1966) observed that the intravenous administration of 0.4-0.8 g/kg ethanol enhanced presynaptic inhibition of primary afferent fibers in the spinal cord of unanesthetized spinal cats. Subsequently, Banna (1969) showed, in cats lightly anesthetized with pentobarbital or decerebrated, that ethanol (3-5 ml of 20% solution/kg i.v.) increased presynaptic inhibition of cutaneous transmission in cuneate and gracile nuclei. Davidson and Rix (1972) then demonstrated that the topical application of ethanol (0.24%, w/v) elicited the manifestations of presynaptic inhibition of primary afferent terminals in the cuneate nucleus. Davidoff (1973) then showed, in in Z J frog ~ spinal cord, that in addition to augmenting presynaptic inhibition, 98 mM ethanol, by bath application, potentiated the action of GABA on primary afferent terminals. In the cortex, Nestoros (1980) recorded extracellularly from neurons in the pericruciate cortex of anesthetized (Fluothane or methoxyflurane) cats using multibarreled microelectrodes and reported that the release of ethanol by electroosmosis potentiated an inhibition of neuronal firing elicited by electrical stimulation of the surface of the cerebral cortex (an effect presumed to be mediated by GABA) and an inhibition of firing produced by the microiontophoretic administration of GABA. He also reported that ethanol did not potentiate the inhibitory action of glycine, serotonin, or dopamine. In these experiments the concentration of ethanol at the membrane of the cell under study was not known, thus Nesteros calculated that the concentration would be 12-120 pM, which is significantly less than the blood-ethanol concentration that gives minimally detectable behavioral effects ( - 5 m). On the other hand, in an earlier study in rat cerebral cortex, Lake et al. (1973) indicated that the electroosmotic release of ethanol did not appear to affect the inhibition of neuronal firing elicited by the microiontophoretic administration of GABA. In the cerebellum, Bloom ut al. (1984), using extracellular recording methods, reported that the intraperitoneal administration of a dose of ethanol calculated to result in a blood-alcohol concentration of 140 mg% did not affect the inhibition of Purkinje cell firing elicited either by an inhibitory synaptic pathway presumed to be mediated by GABA or by the iontophoretic administration of GABA. On the other hand, Harris and Sinclair ( 1984a) also used extracellular recording methods, but found that the local micropressure application of ethanol (750 mM pipette concentration) reduced the inhibition of neuronal firing elicited by a synaptic pathway presumed to be mediated by GABA, and the intravenous administration of 1.5 g/kg ethanol reduced the inhibition produced by the micropressure application of GABA.

A APVll.5 mM Mg2+

Dcd

1 mV

2 msec

Baseline

Wash

B

l 0o -20

* 4

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100

EtOH (mM) FIG.8. Effect of ethanol on pEPSPs mediated by non-NMDA glutamate receptors in hippocampus. (A) Effect of 50 mM ethanol on non-NMDA glutamate receptor-mediated pEPSP. Compare to records in Fig. 6. (B) Average effect of ethanol on non-NMDA glutamate receptor-mediated pEPSPs. Bar graphs show average percent inhibition (mean '. SEM) of non-NMDA glutamate receptor-mediated pEPSPs. The effect of 50 mM ethanol was not statistically significant ( p > 0.1); however the 9% inhibition of non-NMDA glutamate receptor-mediated pEPSPs by 100 m M ethanol was significant (p < 0.025). Recording from rat hippocampal slice. Experimental conditions the same as in Fig. 6 , except external 1.5 mh4 Mg*+, and no 6,7medium contained 50 pkf ~-2-amino-5-phosphonovalerate, dinitro-quinoxaline-2,3-dione. From Lovinger et al. (1990b).

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In the hippocampus, Carlen et al. (1982) recorded intracellularly from CA 1 neurons in the hippocampal slice using an interface chamber and reported that 20 mM ethanol potentiated inhibitory postsynaptic potentials, but did not affect responses to GABA. However, these observations on synaptic inhibition have not been confirmed in subsequent studies on the hippocampus. In an in uzzw study of extracellularly recorded rat hippocampal CA1 and CA3 neurons, Mancillas et al. (1986) concluded that the intraperitoneal administration of ethanol (0.75 or 1.5 g/kg) had no significant effect on the inhibitory response to GABA. Gage and Robertson ( 1985) used the single-electrode voltage-clamp technique to study the CA1 pyramidal neurons in rat hippocampal slices; they found that 10-200 mM ethanol had no perceptible effect on the decay time constant, rise time, or amplitude of inhibitory postsynaptic currents (IPSCs). In intracellular experiments on CA1 and CA3 pyramidal cells in the hippocampal slice, Siggins ef al. (1987) found that 10200 miM ethanol most often reduced the amplitude of IPSPs. In the septal area, Givens and Breese (1990b) found that in medial septal neurons the systemic administration of ethanol ( 1.5 g/kg) potentiated the reduction of firing frequency induced by the iontophoretic application of GABA; however, in lateral septal neurons, ethanol did not affect the GABA-induced reduction of firing frequency. On the other hand, ethanol did not alter the reduction of firing frequency induced by the application of the benzodiazepine flurazepam to medial septal neurons. In addition, electrical stimulation of the fimbria elicited an inhibition of firing frequency in medial septal neurons that was reduced by the application of bicuculline. T h e systemic administration of ethanol prolonged the duration of the reduction of firing frequency induced by fimbrial stimulation. Ethanol (1.5 g/kg) also potentiated the reduction of firing frequency of medial septal neurons induced by nipecotic acid, a GABA uptake inhibitor. T h e observations indicate that ethanol can potentiate GABA responses in the medial septal nucleus and the effect is site specific. Ethanol (10-50 nu2.l) has been reported to elicit a concentrationdependent potentiation of GABA-stimulated "C1- flux in crude brain homogenates (microsacs or synaptoneurosomes) (Allan and Harris, 1986; Suzdak P t al., 1986a, 1988) and cultured spinal cord neurons (Mehta and Ticku, 1988). This potentiation has been reported to be antagonized by the benzodiazepine partial-inverse agonist RO 15-4513 (Suzdak et al., 1986b; Harris et al., 1988; Ticku and Kulkarni, 1988). RO 15-4513 has also been reported to antagonize some of the behavioral effects of ethanol (Bonetti et af., 1985, 1989; Polc, 1985; Suzdak et al., 1986b; Hoffman et al., 1987; Lister, 1987; Koob et al., 1989). However,

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the specificity of RO15-4513 as a selective ethanol antagonist has been questioned in several studies (Bonetti et al., 1984, 1985; Lister and Nutt, 1987; Nutt and Lister, 1987; Britton et al., 1988; Hatch and Jernigan, 1988; Bonetti et al., 1989; Koob et al., 1989).In addition, RO15-4513 has been reported to produce an epileptiform-like pattern of activity in the EEG (Bonetti et al., 1989; Marrosu et al., 1989; Ehlers et al., 1990), to potentiate convulsions (Bonetti et al., 1984, 1989; Miczek and Weerts, 1987; Lister and Nutt, 1988; Ticku and Kulkarni, 1988; Corda et al., 1989),and to activate spike firing in central neurons (Bonetti et al., 1984, 1989; Polc, 1985; Mereu et al., 1987; Marrosu et al., 1989). In electrophysiological studies on the cerebellum, Palmer et al. (1988) showed that the reduction of Purkinje cell firing rate elicited by pressure application of ethanol could be inhibited by the local application of RO15-4513 and FG 7142 (another partial-inverse agonist) from another barrel of the same micropipette. By contrast, RO15-4513 did not antagonize similar depressions of firing induced by local GABA application. Palmer et al. (1990) have also reported that RO15-4513 antagonized the ethanolinduced depression of spike firing in fragments of human fetal cerebellum and cortex transplanted in the anterior eye chamber of athymic nude rats. However, because ethanol did not significantly potentiate GABA responses in the cerebellum, Palmer and Hoffer (1990) concluded that the RO15-4513 inhibition of ethanol responses probably does not occur directly at the GABA, receptor. Clearly, further work is needed on RO 15-4513 to determine the specificity and mechanism of its interaction with ethanol. a. GABA, Channels. There have been several recent voltage-clamp studies of ethanol effects on GABA-activated C1- current, with different results in different preparations. In several studies on neurons in primary tissue culture, potentiation of GABAA current has been reported. Celentano et al. (1988) studied chick spinal cord neurons in tissue culture and reported that when 10-sec pulses of 50 mM ethanol were applied every 3-4 min, a cumulative enhancement of GABA-activated current was observed (often up to double the initial control). Ethanol was applied by pressure ejection from a seven-barrel micropipette. On the other hand, 50 mM ethanol did not affect GABA-activated current with a single prolonged application or with 30-sec pulses. This apparent effect of ethanol did not reverse with wash periods of up to 30 min. The enhancement was observed in 62% of the cells in which current was elicited by 3 pA4 GABA, but no enhancement was observed when current was activated by 30 GABA. In addition, 10-sec pulses of ethanol were reported to have a greater potentiating action on glycine-activated current than on GABA-activated current.

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A CONTROL

H

B

CONTROL

ETHANOL

CONTROL

J FLURAZEPAM

PENTOBARBITAL

:

20 5ec 0 (y

ETHANOL

CONTROL

FIG. 9. Effect of ethanol on GXBA-activated current in cortical neurons. (A) Potentiation b! 20 null etlranol of current activated by 2.5 pbl GABA. (B) Potentiation of current activated by 2.5 wl.M GABA by 20 @ Hurazeparn and 20 p M pentobarbital, hut lack of effect of 40 n&f ethanol. Whole cell patch-clamp recording from cultured mouse cortical netirons. GABA and drugs administered from a macropipette, as in Fig. 1 . From Aguayo ( 1990).

In mammalian neurons, Aguayo (1990) and Aguayo and Weight ( 1990) investigated the effect of ethanol on GABA-activated currents in

hippocampal and cortical neurons cultured from fetal mice. They found that 1-40 null ethanol produced a concentration-dependent potentiation of the current activated by 2.5 ph4 GABA in 70% of the neurons studied (Fig. 9A). In neurons in which ethanol did not potentiate GABAactivated current, this current was still potentiated by flurazepam and pentobarbital (Fig. 9B). When applied in the absence of GABA, ethanol did not induce any membrane current. The ethanol potentiation of GABA-activated current was not voltage dependent at membrane potentials between -40 and +40 mV, and ethanol did not alter the reversal potential of GABA-activated current. Ethanol also potentiated glycineactivated current in these neurons. Nishio and Narahashi (1990) used cultured dorsal root ganglion neurons from neonatal rats to study the effect of ethanol on GABA-activated current. They found that bath application of 30-300 inn.l ethanol produced a concentrationdependent augmentation of the initial transient inward current activated by 100 pfiM GABA, but did not affect the subsequent sustained inward current activated by GABA. On the other hand, ethanol effects on GABA, current have not been observed in voltage-clamp studies on adult mammalian neurons. In neurons freshly dissociated from adult rat dorsal root ganglia, White et al. (1990) investigated the effect of a range of ethanol concentrations on the C1- current activated by a range of GABA concentrations. They found

33 1

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10 pM GABA

1

50 CM GABA

1 Control

A:

20 sec

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FIG. 10. Lack of ethanol effect on GABA-activated current in adult dorsal root ganglion neurons. (Al) Effect of 50 mM ethanol on current activated by 10 pA4 GABA. (A2) Bar graphs showing average (mean k SEM) effect of 10, 50, and 100 mM ethanol on current activated by 10 pA4 GABA. N o group was significantly different from control (p > 0.1; n = number of neurons averaged). (Bl) Effect of 50 mM ethanol on current activated by 50 pA4 GABA. (B2) Bar graphs showing average (mean ? SEM) effect of 10, 50, and 100 mM ethanol on current activated by 50 pA4 GABA. No group was significantly different from control (p > 0.1; n = number of neurons averaged). Whole cell patch-clamp recording from neurons freshly dissociated from adult rat dorsal root ganglion. GABA and ethanol administered from a macropipette, as in Fig. 1. From White et al. (1990).

that ethanol concentrations of 10, 50, and 100 mM did not significantly affect the C1- current activated by 10, 50, or 100 phi GABA (Fig. 10). On the other hand, GABA-activated current was potentiated by pentobarbital and flurazepam in these neurons (White, 1990).Because intracellular Ca2+ and ATP have been reported to affect GABA-activated C1- current, White et al. (1990) also tested the effect of buffering intracellular Ca2 to a very low concentration using a patch-pipette solution containing 10 mM BAPTA and no added Ca2 , as well as 2 mM ATP; +

+

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however, under these conditions 50 mM ethanol had no effect on the current activated by 10 p,M GABA. In addition, because it has been suggested that there are endogenous ligands that interact with benzodiazepine-binding sites (cf. Polc, 1988),White et d.(1990)also tested the effect of ethanol in the presence of both benzodiazepine agonists and inverse agonists. They found that 50 mM ethanol had no significant effect on the current activated by 10 p,M GABA in the presence of either flurazepam or methyl-6,7-dimethoxy-4-ethyl-~-carboline-3-carboxylate (DMCM). In addition, Osmanovic and Shefner (1990)studied the effect of ethanol on GABA-activated current in locus coeruleus neurons in a brain slice preparation. They found that bath application of 40-60 mM ethanol affected neither the inward current nor the conductance increase activated by GABA (200 p,M to 1 mM), although pentobarbital potentiated both parameters. b. Commeiitaq. T h e preceding studies indicate that ethanol can potentiate GABA-activated C1- current in some preparations, but does not do so in others. T h e reason for this difference has not been determined. Some of the factors that may be involved include the type of neuron, the state of cellular elements that regulate GABA, channels (such as G proteins and second messengers, including intracellular Cay and cyclic nucleotides), the state of channel regulation by processes such as phosphorylation and glycosylation, and the types of GABAA subunits expressed in the neuron. In this regard, Wafford el al. (1990)recently reported that ethanol potentiated GABA-activated current in Xenopw oocytes injected with brain mRNA from LS mice, but inhibited GABAactivated current in oocytes injected with mRNA from SS mice. Wafford et ul. ( 1990)also found that 1 p,M RO 15-4513 antagonized the potentiation of GABA current by ethanol in oocytes injected with LS mRNA, but did not affect the inhibition of GABA current by ethanol in oocytes injected with SS mRNA. With respect to the RO15-4513action, it is of interest to note that Luddens et al. (1990)recently reported that recombinant GABA,\ receptors, composed of an a6 subunit in combination with the &,yY subunits, bind with high affinity to RO15-4513,but not the other benmdiazepines or P-carbolines. These observations suggest that genetic factors and the types of channel subunits or their regulation (e.g., Whiting et al., 1990)may be important in the sensitivity of GABA, channels to ethanol. It remains for future research to determine the factors that are responsible for the ethanol sensitivity of GABA, channels. +

3 . Other Trunsmitter-Gated Ion Channels In addition to the excitatory and inhibitory amino acid-activated ion channels, there are also several other transmitter-gated ion channels that

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have been investigated for sensitivity to ethanol. These will be briefly considered below. a. nACh Channels. Ethanol has been found to have effects on synaptic transmission at several cholinergic junctions where the postsynaptic response is mediated by nicotinic acetylcholine (nACh) receptors. In early studies on stellate ganglia of cat, Larrabee and Posternak (1952) found that 1 and 2% ethanol facilitated synaptic transmission, whereas 3% ethanol depressed transmission. I n cat spinal cord, Meyer-Lohmann et al. (1972) demonstrated that the intravenous administration of 0.162.4 g/kg ethanol increased the discharge of Renshaw cells elicited by antidromic stimulation of motor nerve fibers. In early studies at the neuromuscular junction, Feng and Li (1941) found that ethanol facilitated transmission. Subsequent analysis by Gage (1965), Okada (1967), and Quastel and Linder (1975) revealed that the facilitation resulted from both an increase in neurotransmitter release and an increase in the postsynaptic action of the neurotransmitter. Bradley et al. (1980) studied the influence of different concentrations of ethanol on endplate currents elicited by the iontophoretic administration of acetylcholine, and found that 43-430 mm ethanol produced a concentration-dependent potentiation of current amplitude; 43 mM ethanol approximately doubled the amplitude of the ACh current. Gage et al. (1975) and Bradley et al. (1980) used ACh-induced endplate current noise to analyze the mechanism involved in the postsynaptic action of ethanol, and found that ethanol increased the open time of the nACh channel. Because ethanol shifted the concentration-response curve for ACh to the left, Bradley et al. (1980) concluded that ethanol increased the ACh response by decreasing the apparent dissociation constant (Kd) for ACh interaction with its binding site. Nicotinic ACh channels are located in the mammalian CNS, as well as at peripheral cholinergic junctions. Recent molecular biological studies indicate that the nACh channels in the CNS differ somewhat in their amino acid sequences compared to nACh channels at the neuromuscular junction (cf. Deneris et al., 1991; Galzi et al., 1991). Consequently, the sensitivity of central nACh channels to ethanol might differ from that of the neuromuscular junction. Further work is needed to determine the ethanol sensitivity of central nACh channels. b. 5-HT3 Channels. The ethanol sensitivity of the serotonin-activated 5-HT, type of transmitter-gated ion channel has recently been investigated by Lovinger (1991) in NCB-20 neuroblastoma cells using the whole cell patch-clamp technique. He found that ethanol potentiated 5-HT, receptor-mediated current in a concentration-dependent manner at concentrations from 25 to 100 mM (Fig. 11). An ethanol

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A

1 5-HT

5-HT 50 m M EtOH

2

5-HT

150 P A L

lOsec

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8o

1

FIG. 11. Ethanol potentiation of 5-HTs-activated ion current. (A) Effect of 50 mM ethanol on current activated by 1 pM 5-HT. (B) Average percent increase of 5-HTyactivated current as a function of ethanol concentration. Each point in the graph represents t h e mean +- SEM percent potentiation of current activated by 1 pM 5-HT. Whole cell patch-clamp recording from NCB-20 cells. 5 H T and ethanol administered from a macropipette, as in Fig. 1. From Lovinger ( 1 99 1).

concentration of 100 mM increased the amplitude of 5-HT-activated current by 59%. Although ethanol concentrations greater than 100 mh4 also potentiated the current, the potentiation was not greater than that observed with 100 n M ethanol. Potentiation by 50 mM or greater ethanol was observed in 86%of the cells in which the current was elicited by 1 or 2 pA4 5-HT. T h e current activated by the selective 5-HT3 receptor agonist, 2-methyl-5-HT, was also potentiated by 50 o r 100 mM ethanol. Lovinger and White (1991) examined further the potentiation of

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5-HT, current by ethanol in NCB-20 cells and showed that the magnitude of the potentiation decreased with increasing 5-HT concentration. 5-HT, an Thus, 50 mM ethanol produced a 45% potentiation with 1 18% potentiation with 2 p.M 5-HT, and no observed potentiation with 10 5-HT. In addition, the potentiation of 5-HT current was not voltage dependent, the reversal potential of the potentiated 5-HT current was not altered, and the augmented current had an increased decay rate. Lovinger and White (1991) also showed that ethanol potentiated 5-HT, current in neurons freshly dissociated from rat nodose ganglia at concentrations similar to those observed in NCB-20 cells (25- 100 mM). Methanol was less potent than ethanol at potentiating 5-HT3 current. c. ATP Channels. Most recently, the effect of ethanol on the ion channels gated by adenosine 5'-triphosphate (ATP) has been investigated by Li et al. (1991) in neurons freshly dissociated from bullfrog dorsal root ganglia using the whole cell patch-clamp technique. They found that ethanol decreased the amplitude of ATP-activated inward current in a concentration-dependent manner over the range of 6-250 mM. Over this concentration range, ethanol did not elicit membrane current alone, nor did it change the reversal potential of ATP-activated inward current. The average inhibition by 100 mM ethanol of the inward current activated by 5 p.M ATP was 31%. The ethanol concentration that produced 50% inhibition (IC50)of ATP-activated current was 160 mM. Methanol was less potent and 1-propanol was more potent than ethanol in inhibiting ATP-activated current; however, 1-butanol and isopentanol were without effect on this current (C. Li and F. F. Weight, unpublished observations). The percent inhibition of ATP-activated current by ethanol diminished with increasing agonist concentration in a manner consistent with an effect of ethanol on receptor affinity (C. Li and F. F. Weight, unpublished observations). d. Commentary. The preceding studies show that not only are the excitatory and inhibitory amino acid-activated ion channels sensitive to ethanol, but the other currently known neurotransmitter-gated ion channels also show ethanol sensitivity. Although quantitatively the ethanol sensitivity of each type of transmitter-gated ion channel differs, the function of each type of channel is affected in a range of ethanol concentrations that produces behavioral effects. In addition to directly activating ion channels, many neurotransmitters mediate their effects via G proteins and second messengers; however, the effects of ethanol on G proteins and second messengers have received little attention in neurophysiological investigations [for review of neurochemical studies, see Deitrich et al. (1989) and Hoffman and Tabakoff (1990)l.

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V. Summary and Conclusions

Almost a century ago Overton and Meyer observed that the anesthetic potency of different alcohols is correlated with their partition between olive oil and water. This observation led to the “lipid theory of alcohol action,” which has provided a theoretical physicochemical explanation for the mechanism of alcohol action for many years. One of the most recent versions of this theory is the hypothesis that ethanol effects in the nervous system are due to an increase in the fluidity of neuronal membrane lipids. Although criticisms of the lipid theory of alcohol action have been advanced and it has been suggested that alcohols may directly af€ect membrane proteins, the lipid theory of alcohol action has prevailed for many years, in large part because it has not been possible to demonstrate that neuronal membrane proteins are sensitive to alcohol. One approach to elucidating the cellular actions of alcohol in the nervous system has been to investigate the effects of ethanol on the spike firing of neurons in different regions of the nervous system. Such investigations have revealed a plethora of results. Ethanol has been observed to increase, decrease, have biphasic effects on, or have no effect on neuronal discharge in different brain regions. In several brain regions, certain general patterns of ethanol effects have emerged, with either inhibition o r excitation of neuronal firing predominating. On the other hand, in some brain regions, different effects of ethanol administration have been observed on the firing of a given neuronal type dependent upon such factors as the preparation used, the presence or absence of anesthetic, the type of anesthetic used, and the method of ethanol administration. This diversity suggests that a variety of different factors may influence the effect o f ethanol on neuronal discharge. ‘Thus, in order to elucidate the cellular mechanisms of ethanol’s action, it is necessary to minimize the variables that impinge upon neuronal function. The intrinsic electrical excitability of neurons involves voltage-gated membrane ion channels. Investigation of the effects of ethanol on voltage-gated ion channels has revealed that these membrane proteins are relatively insensitive to the effects of ethanol. Although the function of these channels can be inhibited by ethanol, in most cases the concentrations of ethanol needed to significantly inhibit voltage-activated ion currents were well above the blood-ethanol concentrations that are lethal in mammals. Ethanol effects, in pharmacologically relevant concentrations, have been observed on M-current and Ca2+ current in some, but not all, experiments, suggesting that further work is needed to determine whether these differences are due to differences in experimental preparation o r to the sensitivity of these channels to ethanol.

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A. IONCHANNEL HYPOTHESIS OF ALCOHOL ACTION I n contrast to the relative insensitivity of most voltage-gated ion channels to the effects of ethanol, all of the currently known neurotransmittergated ion channels have been reported to be sensitive to ethanol in pharmacologically relevant concentrations. For excitatory amino acidactivated ion channels, the most sensitive channel to the effects of ethanol was the NMDA channel, with a concentration-dependent inhibition of NMDA-activated current over the concentration range 5-50 & ethanol, and IC,, values of 10 and 30 mM for DRG and hippocampal neurons, respectively. Kainate and quisqualate channels were less sensitive to the actions of ethanol, with a threshold for current inhibition of 25 mM and 45% inhibition being observed at 200 mM ethanol. With respect to the inhibitory amino acid-activated ion channels, sensitivity of both GABA- and glycine-gated channels to ethanol has been reported. In contrast to the inhibition of currents activated by excitatory amino acid agonists, ethanol potentiated the currents activated by these inhibitory amino acids. In addition, excitatory amino acid-activated ion channels have been sensitive to ethanol in virtually all neurons tested, whereas GABA-activated channels have been sensitive to ethanol in some neurons and insensitive to ethanol in other neurons. The observation that ethanol has opposite effects on GABA-activated current in oocytes injected with mRNA from LS and SS mice suggests that genetic factors may be important in the ethanol sensitivity of GABA, channels; perhaps differences in the subunits of the GABA, channel that are expressed or their regulation are determinants of channel sensitivity to ethanol. In this regard, it is of interest to note that the current associated with GABAA channels formed from a and (3 subunits is not potentiated by benzodiazepines; the presence of the y subunit is necessary for potentiation (Pritchett et al., 1989). In addition to the excitatory and inhibitory amino acid-activated ion channels, other neurotransmitter-gated ion channels have also been found to be sensitive to the actions of ethanol. For both nACh channels and 5-HT3 channels, the transmitter-activated current was potentiated by ethanol. An ethanol concentration of 43 mM approximately doubled the amplitude of nACh current, with increasing concentrations producing increasing augmentation of current amplitude. 5-HT3 current was potentiated in a concentration-dependent manner by ethanol concentrations from 25 to 100 mM, with 100 mM ethanol increasing the current by 59%. ATP channels were also found to be sensitive to ethanol; however, in the case of these channels, ethanol inhibited ATP-activated current. This current was decreased by ethanol in a concentrationdependent manner over the range 6-250 mM, with an IC,, of 160 &.

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The mechanism of ethanol effects on transmitter-gated ion channels is not well understood and appears to differ at different channels. For both 5-HT, and ATP channels, increasing agonist concentration diminished the effect of ethanol on channel current, an action consistent with an action on the receptor, perhaps altering its affinity for agonist. However, because ethanol potentiated 5-HT, current but inhibited ATP current, this interpretation would suggest that ethanol may increase the affinity of the 5-HT, receptor and decrease the affinity of the ATP receptor. On the other hand, the inhibition of NMDA current by ethanol did not differ with different agonist concentrations. In addition, ethanol did not appear to reduce NMDA current by a channel-blocking action, by altering the ion selectivity of the channel, or by affecting several regulatory sites on the channel. Taken together with the data that the potency of several alcohols for inhibiting NMDA current is correlated with the hydrophobicity of the alcohols, these observations are consistent with the idea that ethanol may inhibit NMDA current by an interaction with a hydrophobic site associated with the NMDA channel. In this context, it is of interest to note that although methanol, ethanol, and 1propanol exhibited increasing potency for inhibiting ATP current, 1butanol and isopentanol had no effect on this current. This observation is difficult to explain in terms of the lipid hypothesis, because 1-butanol and isopentanol are more hydrophobic than methanol, ethanol, and 1propanol. In addition, this observation suggests that the active site of alcohol action associated with the ATP channel may be a small hydrophobic pocket or there may be limited access to the hydrophobic site, for example, via the channel pore. T h e preceding observations are consistent with the hypothesis that transmitter-gated ion channels are ethanol-sensitive membrane proteins. Molecular biologists consider the hydrophobic regions of channel proteins to be the membrane-spanning domains. This implies that if ethanol interacts with a hydrophobic site on the channel protein, the 'most likely site of ethanol's interaction would be the membrane-spanning domain. T h e hypothesis that ethanol interacts with a hydrophobic region of transmitter-gated ion channels has considerable heuristic value, because it suggests experiments to ascertain the molecular basis of ethanol's action. For example, if an ethanol-sensitive channel is cloned and sequenced, site-directed mutagenesis could be used to determine the ethanol-sensitive region of the channel protein. Also, for channel proteins with ethanol-sensitive and ethanol-insensitive forms, such as the GABA, channel, different combinations of subunits could be expressed in oocytes to determine whether the ethanol sensitivity resides in a particular subunit o r combination of subunits o r whether a regulatory pro-

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cess such as phosphorylation or glycosylation of a particular subunit might confer ethanol sensitivity on the channel.

B. ALCOHOL SENSITIVITY OF TRANSMITTER-GATED IONCHANNELS AND BEHAVIORAL EFFECTS OF ALCOHOL T h e observations that transmitter-gated ion channels are sensitive to ethanol concentrations that produce behavioral effects in animals raise the question of whether the actions of ethanol on these ion channels are responsible for the behavioral effects of ethanol. In regard to this question, the range over which ethanol produced concentration-dependent inhibition of NMDA-activated current corresponds to the range over which intoxication is observed in nontolerant humans. Impairment of fine motor control and delayed reaction time can be detected at bloodethanol concentrations of 20-30 mg%, and increasing blood-ethanol concentration to 200-250 mg% results in increased impairment of motor coordination and cognitive function, which are recognized as signs of intoxication (Ritchie, 1985;Kissin, 1988).In this regard, it is of interest to note that the intracerebroventricular administration of the specific NMDA receptor antagonist, D-2-amino-5-phosphonovalerate (APV), produces behavioral effects in animals similar to those observed in ethanol intoxitation (Woods et al., 1987).These observations suggest that ethanol inhibition of responses to NMDA receptor activation may contribute to the neural and cognitive impairments associated with intoxication. I n contrast to the effect of ethanol on NMDA-activated current, ethanol inhibition of kainate- and quisqualate-activated currents increased significantly at concentrations greater than 50 mM, which corresponds to the concentration range of ethanol that produces general anesthesia. Clinically, blood-ethanol concentrations greater than 200-250 mg% are associated with progressive CNS depression and behavioral signs of general anesthesia, characterized by increasing sedation, which progresses to stupor and coma; death from respiratory depression begins to occur at concentrations greater than 500 mg% (Kissin, 1988). Because glutamate is thought to be the major excitatory transmitter in the CNS, and kainate and quisqualate receptors mediate fast synaptic transmission at glutamatergic synapses, inhibition of kainate and quisqualate receptoractivated responses by ethanol would be expected to result in general CNS depression. In support of this concept, the anesthetic barbiturates inhibit kainate- and quisqualate-activated currents more potently than NMDA-activated current (Peoples et al., 1990;Weight et al., 1991),and

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trichloroethanol, the active metabolite of the general anesthetic chloral hydrate, inhibits kainate- and quisqualate-activated current more potently than ethanol (Peoples and Weight, 1991b). These observations suggest that ethanol inhibition of responses to kainate and quisqualate receptor activation may contribute to the general anesthetic effect of ethanol. In most studies in which ethanol affected GABAA responses, ethanol potentiation of GABA, responses did not increase with ethanol concentrations greater than 50 mM. this would suggest that the augmentation of GABA, current by ethanol does not contribute significantly to the general anesthetic effects of ethanol, because these effects occur at blood-ethanol concentrations greater than 200-250 mg%. On the other hand, in these studies, ethanol produced a concentration-dependent potentiation of GABA, responses over the range of 10-50 mM. Because this is similar to the concentration range that produces intoxication, it raises the question of whether the potentiation of GABAA responses by ethanol contributes to the intoxicating actions of ethanol. A number of behavioral studies have suggested that GABAA responses are involved in the intoxicating actions of ethanol. This is supported by the observation that RO15-4513 blocks some of the intoxicating actions of ethanol and the ethanol potentiation of 36Cl- flux in brain homogenates and cultured spinal neurons. On the other hand, benzodiazepines also potentiate GABA, responses, but d o not produce an intoxicated state in their clinical dose range. Benzodiazepines are widely used clinically, primarily for their anxiolytic actions (cf. Baldessarini, 1985), suggesting that the potentiation of GABA, responses by ethanol may contribute to the anxiolytic actions of ethanol. There has been considerable recent interest in 5-HT, channels based on reports that selective 5-HT, antagonists can block the rewarding properties of morphine and nicotine (Carboni et al., 1989a). The actions of these drugs, as well as of ethanol, have been associated with the release of dopamine in the nucleus accumbens (Di Chiara and Imperato, 1988), and 5-HT, antagonists have been reported to block morphine-, nicotine-, and ethanol-induced release of dopamine from the nucleus accumbens (Carboni et al., 1989b; Wozniak et al., 1990). In behavioral experiments, Grant and Barrett (1991) have reported that the 5-HT3 antagonists ICS 205-930 and MDL 72222 can block the ability of pigeons to discriminate between ethanol and water. This suggests that ethanol potentiation of 5-HT, current may contribute to the ability to discriminate the action of ethanol. Whether 5-HT, channels are also involved in the rewarding properties of ethanol remains to be determined in future investigations.

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The behavioral significance of the other ethanol-sensitive transmittergated ion channels is not known. The role that nACh channels in the CNS play in behavior has not been determined. Presumably, the behavioral and rewarding effects of nicotine involve nACh receptors. However, whether or not the potentiating action of ethanol on nACh current contributes to behavior remains to be determined. Glycine is believed to be the major inhibitory neurotransmitter in the spinal cord (Curtis and Johnston, 1974). Thus, the potentiation of glycine-activated current by ethanol that has been observed in some experiments would be expected to augment IPSPs in the spinal cord, and thereby modulate spinal reflex activity. However, in a study of the spinal cord, ethanol potentiation of spinal IPSPs was not apparent (Klee et al., 1975), raising the question of whether there may be ethanol-sensitive and ethanol-insensitive forms of glycine-gated channels, as there appears to be for GABA-gated channels. ATP-gated ion channels have been demonstrated in several peripheral excitable cells (cf. Bean and Friel, 1989) and sensory neurons (Bean, 1990), but the distribution and function of ATP-gated ion channels in the CNS is not yet known. Consequently, the functional significance of ethanol inhibition of ATP-activated ion current remains to be determined. Further work is clearly needed to elucidate the action of ethanol at these transmitter-gated ion channels in the CNS, and the behavioral significance of these actions.

C. CONCLUSIONS Research investigations beginning almost a century ago have shown that the intoxicating and anesthetic properties of alcohols are correlated with their oillwater or membranelbuffer partition coefficients. These observations led to what has been called the “lipid theory of alcohol action.” More recent investigations on the effects of ethanol on neuronal discharge in the CNS indicate that ethanol can produce a number of effects on spike firing. The effects of ethanol on action potential discharge may be influenced by a variety of different variables, making interpretation of the cellular mechanisms of ethanol action difficult. Investigations on the cellular mechanisms of ethanol action using in vitro preparations have shown that voltage-gated ion channels are relatively resistant to the effects of ethanol, with most effects being observed at ethanol concentrations that would be lethal in mammals. On the other hand, all of the currently known transmitter-gated ion channels have been reported to be sensitive to ethanol in pharmacologically relevant concentrations. The ethanol sensitivity of transmitter-gated ion channels

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may result from an interaction of ethanol with a hydrophobic domain of these membrane proteins. This concept has been designated the “ion channel hypothesis of alcohol action.” T h e sensitivity of different transmitter-gated ion channels to ethanol appears to explain many of the behavioral effects of ethanol. It is suggested that the ion channel hypothesis of alcohol action will be a useful theoretical framework for elucidating the molecular basis of alcohol action. Moreover, the ion channel hypothesis should provide a basis for investigating the pathophysiologic mechanisms involved in alcohol abuse and alcoholism.

References

Aguayo, L. G. (1990). Eur. J. Phurmacol. 187, 127-130. Aguayo, L. G., and Weight, F. F. (1988).J. Physiol. (London) 405, 397-419. Aguayo, L. G., and Weight, F. F. (1990). FASEBJ. 4, A741. Alkdn, A. M., and Harris, R. A. (1986). Life Sci. 39, 2005-2015. Arnistrong, C. M., and Binstock, L. (1964).J. Cen. Phy~iol.48, 265-277. Aston-Jones, G., and Bloom, F. E. (1981).J. Neurosci. 1, 887-900. Aston-Jones, G . , Foote, S. L., and Bloom, F. E. (1982). Nature (London) 296, 857-860. Augustine, G. J., Charlton, M. P., and Smith, S. J. (1987).Annu. Rev. Neurosci. 10,633-693. Baldessarini, R. J. (1985). In “The Pharmacological Basis of Therapeutics” (A. G. Gilman, L. S. Goodman, T. W. Rall, and F. Murad, eds.), 7th ed., pp. 433-438. Macmillan, New York. Banna, N. R. (196Y). Experzentiu 25,619-620. Bade, A., Hofter, B., and Dunwiddie, T. (1983). Brain Res. 264, 69-78. Bean, B. P. (1990).J. Neurosci. 10, 1-10, Bean, B. P., and Friel, D. D. (1989). In ”Ion Channels,” (T.Narahashi, ed.), Vol. 2, pp. 169203. Plenum, New York. Benron, D. M., Blitzer, R. D., and Landau, E. M. (1989). Eur. J. Phrmarol. 164,591-594. Berger, T., French, E. D., Siggins, G. R.,Shier, W. T., and Bloom, F. E. (1982). Pharmucol., Biochem. Behat. 17, 8 13-82 1. Bergmann, M. C . , Klee, M. R.,and Faber, D. S. (1974). Pfiuegers Arch. 348, 139-153. Bloom, F. E., Siggins, G. R., Foote, S. L., Gruol, D., Aston-Jones, G., Rogers,J., Pittman, Q., and Staunton, D. (1984). In “Catecholamines: Neuropharmacology and Central Nervous System: Theoretical Aspects” (E. Usdin et d., eds.), pp. 159-167. Liss, New York. Boggs, J. M., Yoong, T., and Hsia, J. C. (1976). Mol. Plurmacol. 12, 127-135. Bonetti, E. P.,Polc, P., and Pieri, L. (1984). Neurosci. t e l t . 18, Suppl., S30. Bonetti, E. P., Burkard, W. P., Gabl, M.,and Mohler, H. (1965). Br.J. P h n m c o l . 86,463P. Bonetti, E. P. et al. (1989). Phunna~ol.,Biochem. Behuv. 31, 733-749. Borg. S., Kvande, H., Mossberg, D., Valverius, P., and Sedvall, G. (1983). Pharmacol., Bzochem. Behnv. 18, Suppl. 1, 37.5-378. Bradley, R. J., Peper, K., and Sterz, R. (1980). Nature (London) 284, 60-62. Bradley, R. J.. Stertz, R., and Peper, K. (1984). Brain Res. 295, 101-1 12. Breese, G. R.,Givens, B. S., McCown, T. J., and Criswell, H. E. (1988). i n “Biomedicat and Social Aspects of Alcohol and Alcoholism” (K. Kuriyma, A. Takada, and H. Ishii, eds.), pp. 273-276. Elsevier, Amsterdam.

ALCOHOL ACTIONS I N THE NERVOUS SYSTEM

343

Brehm, P., and Eckert, R. (1978). Science 202, 1203-1206. Britton, K. T., Ehlers, C. L., and Koob, G. F. (1988). Science 239, 648-649. Brodie, M. S., Shefner, S. A,, and Dunwiddie, T. V. (1990). Brain Res. 508, 65-69. Brown, D. (1988). T r e d Neurosci. 11, 294-299. Buck, K. J., Allan, A. M., and Harris, R. A. (1989). Eur. J. Phurmacol. 160, 359-367. Camacho-Nasi, P., and Treistman, S. N. (1986). Cell Mol. Neurobiol. 6, 263-279. Camacho-Nasi, P., and Treistman, S. N. (1987). Cell Mol. Neurobiol. 7, 191-207. Carboni, E., Acquas, E., Leone, P., and Di Chiara, G. (1989a). Psychopharmacology (Berlin) 97, 175-178. Carboni, E., Acquas, E., Frau, F., and Di Chiara, G. (1989b). Eur.J. Phrmacol. 164,515519. Carlen, P. L., Gurevich, N., and Durand, D. (1982). Science 215, 306-309. Carlen, P. L., Zhang, L., and Cullen, N. (1991). Ann. N.Y. Acud. Sci. 625, 17-25. Celentano, J. J., Gibbs, T. T., and Farb, D. H. (1988). Brain Res. 455, 377-380. Chin, J. H., and Goldstein, D. B. (1977). Mol. Phurmacot. 13,435-441. Chu, N.-S. (1983). 1nt.J. Neurosci. 21, 265-278. Chu, N.-S. (1984). Brain Res. 311, 348-353. Chu, N.-S., and Keenan, L. (1987). Alcohol 4, 373-374. Cline, H. T., Debski, E., and Constantine-Paton, M. (1987).Proc. Nutl. Acad. Sci. U.S.A. 84, 4342-4345. Cole, W. H., and Allison, J. B. (1930).]. Gen. Physiol. 14, 71-86. Collingridge, G. L., and Lester, R. A. J. (1989). Pharmacol. Reu. 41, 143-210. Collins, J. G., and Roppolo, J. R. (1980).J. Pharmucol. Ex#. Ther. 213, 337-345. Corda, M. G., Giorgi, O., Longoni, B., and Biggio, G. (1989). Eur.J. Pharmacol. 159,233239. Curtis, D. R., and Johnston, G. A. R. (1974). Ergeb. Physiol., Biol. Chem. Exp. Pharmukol. 69, 97-188. Daniell, L. C., and Harris, R. A. (1989).J. Pharmacol. Exp. Ther. 250, 875-881. Daniell, L. C., Brass, E. P., and Harris, R. A. (1987). Mol. Phamacol. 32, 831-837. Davidoff, R. A. (1973). Arch. Neurol. (Chicago) 28, 60-63. Davidson, M., Wilce, P., and Shanley, G. (1988). Neurosci. Lett. 89, 165-169. Davidson, N., and Rix, K. J. B. (1972). J. Physiol. (London) 227, 24P-26P. Deitrich, R. A., Dunwiddie, T. V.,Harris, R. A., and Erwin, V. G. (1989).Ph-ol. Rev. 41, 489-537. Deneris, E. S., Connolly,J., Rogers, S. W., and Duvoisin, R. (199 1). Trends Phannncol. Sci. 12, 34-40. Di Chiara, G., and Imperato, A. (1988). Proc. Natl. Acad. Sci. U.S.A. 85, 5274-5278. Dildy, J. E., and Leslie, S. W. (1989). Brain Res. 499, 383-387. Dingledine, R., Hynes, M. A,, and King, G. L. (1986).J. Physiol. (London) 380, 175-189. Doller, H. J., Eckardt, M. J., and Weight, F. F. (1980). Fed. Proc., Fed. Am. SOC.Exp. Biol. 39, 281. Ehlers, C. L., Chaplin, R. I., and Koob, G . F. (1990). Pharmacol., Biochem. Behau. 36,607611. Eidelberg, E., and Wooley, D. F. (1970). Arch. Int. Phurmucodyn. Ther. 185, 388-396. Eidelberg, E., Bond, M. L., and Kelter, A. (1971). Arch. Int. Phurmacodyn. Ther. 192, 213219. Eskuri, S. A,, and Pozos, R. S. (1987). Alcohol Drug Res. 7, 153-162. Feng, T. P., and Li, T. H. (1941). Chin.J. Physiol. 16, 317-340. Ferguson, J. (1939). Proc. R. Soc. London, Ser. B 127, 387-404. Fibiger, H. C., Le Piane, F. G., Jakubovic, A,, and Phillips, G. (1986).J. Neurosci. 7, 38883896.

344

FORREST F. WEIGHT

Forney, E., and Klemm, W. R. (1976). Res. Commun. Chem. Pathol. Phannacol. 15,801-804. Franklin, C . L., and Gruol, D. L. (1987). Brain Res. 416,205-218. Franks, N. P., and Lieb, W. R. (1978). Nature (London) 274, 339-342. Franks, N. P., and Lieb, W. R. (1979). J. Mol. Biol. 133, 469-500. Franks, N. P., and Lieb, W. R. (1982). N d u r e (London) 300,487-493. Franks, N. P., and Lieb, W. R. (1984). Nature (London) 310, 599-601. Franks, N. P., and Lieb, W. R. (1985).Nature (London) 316, 349-351. Franks, N. P., and Lieb, W. R. (1986). Proc. Nafl. Acad. Sci. U.S.A. 83, 5116-5120. Franks, N. P., and Lieb, W. R. (1987). Trends Phannacol. Sci. 8, 169-174. Gage, P. W. (1965). J. Pharmacol. Exp. Ther. 150, 236-243. <;age, P. W., and Robertson, B. (1985). Br. J. Phannacol. 85, 675-681. Gage, P. W., McBurney, R. N., and Schneider, G. T. (1975).J . Physiol. (London) 244, 409429. Galzi, J.-L., Revah, F., Bessis, A., and Changeux, J.-P. (1991). Annu. Rev. Pharmacol. Toxicol. 31, 37-72. Celler, I. (1973). Hiol. Biochem. Behau. 1, 361-365. George, F., and Chu, N.-S. (1984). Alcohol 1, 353-358. Gessa, G . L., Muntoni, F., Collu, M., Vargiu, L., and Mereu, G. (1985).BrainRes. 348,201203. Gill, K., and Amit, 2. (1989). Recent Dev. Akohol. 7, 225-248. Givens, B. S., and Breese, G. R. (1990a). J. Phannacol. Exp. Thw. 253,95-103. Givens, B. S., and Breese, G. R. (1990b). J. Pha-01. Exp. Ther. 254, 528-538. Goldstein, D. B. (1984). Annu. Rev. Pharmacol. Toxuol. 24, 43-64. Gonzales, R. A., and Woodward, J. J. (1990). J . Phannacol. Exp. Ther. 253, 1138-1144. Gothert, M., and Fink, K. (1989).Naunyn-Schmiedeberg’s Arch. Pharmacol. 340, 516-521. Grant, K. A., and Barrett, J. E. (1991). Psychopharmacology (Berlin) 104,451 - 456. Gruol, D. L.. (1982). Brain Res. 243, 25-33. Crupp, L. A. (1980). Psychophannacobgy (Berlin)70, 95-103. Grupp, L. A., and Perlanski, E. (1979). Nnrropharmacology 18, 63-70. Gustafsson, B., and Wigstrom, H. (1988). Trendr Neurosci. 11, 156-162. Harris, D. P., and Sinclair, J. G. (1984a). Ga.Phannacol. 15, 449-454. Harris, D. P., and Sinclair, J. G. (1984b). Gen. Phannacol. 15, 455-459. Harris, R. A., Allan, A. M., Daniell, L. C., and Nixon, C . (1988).J. Phannacol. Exp. Thw. 247, 1012-1017. Hatch, R. C., and Jernigan, A. D. (1988). Life Sci. 42, 11-19. Hill, S. Y. (1974). B b l . Psychiatry 8, 151-158. Hille, B. (1984). “Ionic Channels of Excitable Membranes.” Sinauer Assoc., Sunderland, Massachusetts. Hoffman, P. L., and Tabakoff, B. (1990). FASEBJ. 4,2612-2622. Hoffman, P. L., Tabakoff, B., Szabo, G . , Suzdak, P., and Paul, S. (1987). Life Sci. 41,611619. Hoffman, P. L., Rabe, C. S., Moses, F., and Tabakoff, B. (1989).J. Neurochem. 52, 19371940. Houck, D. J. (1969). Am. J. Physiol. 216, 364-367. Ikeda, S. R., Schofield, G. G., and Weight, F. F. (1986). J.Neurophysiol. 55, 527-539. 325, 529-531. Johnson, J. W., and Ascher, P. (1987). Nature (Lo&) Johnson, S. W., Hoffer, B. J., Baker, R., and Freedman, R. (1985). Alcohol.: Clin. Exp. Res. 9, 56-58. Kissin, B. (1988). I n ‘Cecil Textbook of Medicine” (J. B. Wyngaarden and I. H. Smith, eds.), pp. 48-52. S u n d e r s , Philadelphia, Pennsylvania.

ALCOHOL ACTIONS IN THE NERVOUS SYSTEM

345

Kleckner, N. W., and Dingledine, R. (1988).Science 241, 835-837. Klee, M. R.,Lee, K. C., and Park, M. R. (1975).Pfluegers Arch. 355, R85. Kleinschmidt, A., Bear, M. F., and Singer, W. (1987).Nature (London) 238, 355-358. Koob, G. F.,Percy, L., and Britton, K. T. (1989).Pharmacol., Biochern. Behav. 31, 757-760. Kuba, K. (1980).J.Physiol. (London) 298, 251-269. Lake, N., Yarbrough, G. G., and Phillis, J. W. (1973).J. Pharm. P h a m o l . 25, 582-584. Larrabee, M. G., and Posternak, J. M. (1952).J.Neurophysiol. 15, 92-114. Lewis, D. L., and Weight, F. F. (1988).Neurooendocrinology 47, 169-175. Lewis, D. L., Weight, F. F., and Luini, A. (1986).Proc. Natl. Acad. Sci. U.S.A. 83,9035-9039. Li, C., Aguayo, L., and Weight, F. F. (1991).IBRO World Congr. Neurosci. Abstr., ?rd, 64. Lieb, W. R., Kovalycsik, M., and Mendelsohn, R. (1982).Biochim. Biophys. Acta 688, 388398. Lima-Landman, M. T., and Albuquerque, E. X. (1989).FEBS Lett. 247,61-67. Linnoila, M., Mefford, J., Nutt, D., and Adinoff, B. (1987).Ann. Inten. Med. 107,875-889. Lister, R. G.(1987).Life Sci. 41, 1481-1489. Lister, R. G., and Nutt, D. J. (1987).Trends Neurosci. 10, 223-225. Lister, R. G., and Nutt, D. J. (1988).Br.1. Pharmucol. 93,210-214. Llinas, R. R. (1988).Science 242, 1654-1664. Lovinger, D. M. (1991).Neurosci. Lett. 122,57-60. Lovinger, D. M., and Weight, F. F. (1988).Int. Symp. Alc. Seizures Abstr., p. 24. Lovinger, D. M.,and White, G. (1991).Mal. Pharmacol. (in press). Lovinger, D. M., White, G., and Weight, F. F. (1989).Science 243, 1721-1724. Lovinger, D. M., White, G., and Weight, F. F. (1990a).Ann. Med. 22, 247-252. Lovinger, D. M., White, G., and Weight, F. F. (1990b).J.Neurosci. 10, 1372-1379. Luddens, H., Pritchett, D. B., Kohler, M., Killisch, I., Keinanen, K., Monyer, H., Sprengel, R.,and Seeburg, P. H. (1990).Nature (London) 346, 648-651. Lyon, R. C., McComb, J. A., Schreurs, J., and Goldstein, D. B. (1981).J. Phannacol. Exp. Thm. 218,669-675. MacDermott, A. B., and Dale, N. (1987).Trends Neurosci. 10, 280-284. MacDermott, A. B., Mayer, M. L., Westbrook, G. L., Smith, S. J., and Barker, J. L. (1986). Nature (London) 321, 519-522. Machu, T., Woodward, J. J., and Leslie, S . W. (1989).Alcohol 6,431-436. Mancillas, J. R.,Siggins, G.R.,and Bloom, F. E. (1986).Science 231, 161-163. Marrosu, F.,Carcangiu, G., Passino, N., Aramo, S., and Mereu, G. (1989).Synapse 3, 117128. Mayer, M. L., and Westbrook, G. L. (1987).Prog. Neurobiol. 28, 197-276. Mayer, M. L., Westbrook, G. L., and Guthrie, P. B. (1984).Nature (London) 309, 261-263. McCreery, M.J., and Hunt, W. A, (1978).Neurophamology 17,451-461. McQuilkin, S . J., and Harris, R. A. (1990).Life Sci. 46,527-541. Mehta, A. K., and Ticku, M. K. (1988). J. P h a m o l . Exp. Ther. 246,558-564. Mereu, G., and Gessa, G. L. (1985).Brain Res. 360, 325-330. Mereu, G., Fadda, F., and Gessa, G. L. (1984).Brain Res. 292, 63-69. Mereu, G., Passino, N., Carcongin, P., Boi, V., and Gessa, G. L. (1987).Eur. J. Phannacol. 135, 453-454. Meyer, H. (1899).Arch. Exp. Pathol. P h a m h o l . 42, 109-118. Meyer, H. (1901).Arch. Exp. Pathol. Phannakol. 46,338-346. Meyer, K. H.(1937).Trans. Farwhy SOC.33, 1062-1068. Meyer-Lohmann, J., Hagenah, R., Hellweg, C., and Benecke, R. (1972).NaunynSchmiedeberg’s Arch. Phannacol. 272, 131- 142.

346

FORREST F. WEIGH?’

Meyers, M. D., and Martin, G. E. (1973). Ann. N. Y. A c d . Sci. 215, 135-144. Meyers, R. D., and Melchior, G. L. (1975). Res. Cummutz. Chem. Pathol. Phunnacol. 10,363377. Miczek, K. A., and Weerts, E. M. (1987). Science 235, 1127. Mikeska,J. A., and Klemrn, W. R. (1979). Res. Commun. Psychol. Psychiatry Behuv. 4,457-466. Miyahara, J. T., Esplin, D. W., and Zablocka, B. (1966).J. Phunnacol. Exp. Ther. 154, 119127. Moore, J. W. (1966). Psychom. M o d . 28,450-457. Moore, J. W., Ulbricht, W., and Takata, M. (1964). J. Gen. Physiol. 48, 279-295. Moore, S. D., Madamba, S. G., Joels, M., and Siggins, G. R. (1988). Science 230, 278-280. Moore, S. D., Madaniba, S. G., and Siggins, G . R. (1990). Brain Res. 516, 222-228. IMorris, R. G. M.,Anderson, E., Lynch, G. S., and Baudry, M. (1984). Nature (London) 319, 774-776. Mullins. L. J . (1954). Chein. Rev. 54, 289-323. Mullins. L. J., and Gaffey C . T. (1954). Proc. Soc. Exp. Biol. M e d . 85, 144-149. Naranjo, C. A,, and Sellers, E. M. (1989). Recent Dav. Alcohol 7, 255-266. Nestoros, J. N. (1980). Science 209, 708-710. Niesen, C. E., Baskys, A,, and Carlen, P. L. (1988). Brain Res. 445, 137-141. Nishio, M..and Narahashi, T. (1990). Brain Res. 518, 283-286. Nowak, L., Bregestovski, P., Ascher, P., Herbert, A., and Prochaintz, A. (1984). Nature (London) 307, 462-465. Nutt, D. J., and Lister, R. G. (1987). Brain Res. 413, 193-196. Oakes, S. G., and Pozos, R. S. (1982a). Dev. Brain Res. 5, 243-249. Oakes, S. G., and Pozos, R. S . (1982b). Dev. Brain Res. 5, 251-255. Okada, K. (1967). Jpn. .I. Physiol. 17, 245-261. Osmanovic, S. S., and Shefner, S. A. (1990). Brain Res. 517, 324-329. Overton, E. (1896). Z . Phys. Chm. 22, 189-209. Overton, E. (1901). “Studien uber die Narkose.” Fischer, Jena. Oyama, Y., Akaike, N., Nishi, K. (1986). Brain Res. 376, 280-284. Palmer, M. R.,and Hoffer, B. J. (1990). Neurochern. Res. 15, 145-151. Palmer, hl. R., Sorenson, S. M., Freedman, R., Olsen, L.. Hoffer, B., and Seiger, A. (1982). J. Phaanlmacol. Exp. Ther. 222, 480-487. Palmer, M. R., Wang, Y., Fossom, L. H., and Spuhler, K. P. (1987). Alcohol.: Clin. Exp. Ras. 11,494-501. Palmer, M. R., van Norne, C., Harlan, J. T., and Moore, E. A. (1988). J. Pharmacol. Exp. Tho. 247, 10 18- 1024. Palmer, M. K., Eriksdotter-Nilsson, M., Bygdeman, M., Stieg, P., Stromberg, I., Olsen, L., Seiger, A., Hoffer, B. J . , and Granholm, A. C. (1990). J . Phunnacol. Ex#. Ther. 254, 1100-1 106. Pdnde, U., Pant, H. C., and Weight, F. F. (1983). Soc. Neurosci. Abstr. 9, 1235. Patel, S., Chapman, A. G., hfillan. M. H., and Meldrum, B. S. (1988). In “Excitatory Amino Acids in Health and Disease” (D. Lodge, ed.), pp. 358-378. Wiley (Interscience),New York. Pennefather, P., and Quastel, D. M. J. (1980). I n ”Molecular Mechanisms of Anesthesia” (B. R. Fink, ed.). pp. 45-57. Raven Press, New York. Peoples, R. W., and Weight, F. F. (1991a). Bruin Res. (in press). Peoples, R. W., and Weight. F. F. (199lb). IBRO World Congr. Neurosci. Abstr., 3rd, 63. Peoples. R. W., Lovinger, D. M., and Weight, F. F. (1990). Soc. Neurosci. Abstr. 16, 1017. Phillips, A. G., and Fibiger, H. C. (1978). Can. J . Physiol. Phurmacol. 32, 58-66. Pitler, T. A , , and Landfield, P. W. (1987). Brain Res. 410, 147-153.

ALCOHOL ACTIONS IN THE NERVOUS SYSTEM

347

Pohorecky, L. A., and Brick, J. (1977). Brain Res. 131, 174-179. Polc, P. (1985). Br. J. Phannacol. 86, 465. Polc, P. (1988). Prog. Neurobiol. 31, 349-423. Pritchett, D. B., Sontheimer, H., Shivers, B. D., Ymer, S. Kettenmann, H., Schofield, P. R., and Seeburg, P. H. (1989). Nature (London) 338, 582-585. Quastel, D. M. J., and Linder, T. M. (1975). In “Molecular Mechanisms of Anesthetics” (B. R. Fink, ed.), pp. 157-168. Raven Press, New York. Quevedo, L., Baldeig, J., and Concha, J. (1976). Pharmacology 14, 148-152. Rabe, C. S., and Tabakoff, B. (1990). Mol. Pharmacol. 38, 753-757. Rabe, C. S., and Weight, F. F. (1988).J. Phannacol. Exp. Ther. 244, 417-422. Rang, H. P. (1960). Br. J. Phannacol. 15, 185-200. Rauschecker, J. P., and Hahn S. (1987). Nature (London) 238, 183-185. Rezazadeh, S. M., Woodward, J. J., and Leslie, S. W. (1989). Alcohol 6, 341-345. Ritchie, J. M. (1985).In “The Pharmacological Basis of Therapeutics” (A. G. Goodman, L. S. Gilman, T. W. Rall, and F. Murad, eds.), 7th ed., pp. 372-373. Macmillan, New York. Rogers, J., Siggins, G. R., Schulman, J. A,, and Bloom, F. E. (1980). Brain Res. 196, 183198. Rogers, J., Madamba, S. G., Staunton, D. A., and Siggins, G. R. (1986). Brain Res. 385,253262. Rothman, S. M., and Olney, J. W. (1987). Trends Neurosci. 10, 299-302. Rudy, B. (1988). Neuroscience 25, 729-749. Schwartz, M. H. (1985). Bruin Res. 332, 337-353. Scott, R. H., Wooton, J. F., and Dolphin, A. C. (1990). Neuroscience 38, 285-294. Seeman, P. (1972). Pharmacol. Rev. 24, 583-655. Seil, F. J., Leiman, A. L., Herman, M. M., and Fisk, R. A. (1977).Exp. Neural. 55,390-404. Shah, J., and Pant, H. C. (1988). Brain Res. 474, 94-99. Shefner, S. A., and Osmanovic, M. E. (1988). Alcohol.: Clin. Exp. Res. 12, 323. Shefner, S. A., and Tabakoff, B. (1985). Alcohol 2, 239-243. Siggins, G. R., and French, E. (1979). Drug Alcohol Depend. 4, 239-243. Siggins, G. R., Pittman, Q. J., and French, E. D. (1987).Bruin Res. 414, 22-34. Simson, P. E., Criswell, H. E., Johnson, K. B., Hicks, R. E., and Breese, G. R. (1991).J. Phurmacol. Exp. Ther. 257, 225-23 1 . Sinclair, J. G., and Lo, G. F. (1981). Bruin Res. 204, 465-471. Sinclair, J. G., Lo, G. F., and Tien, A. F. (1980). Can. J. Physzol. Pharmacol. 58, 429-432. Sorenson, S., Palmer, M., Dunwiddie, T., and Hoffer, B. (1980). Science 210, 1143-1 145. Sorenson, S., Carter, D., Marwaha, J., Baker, R., and Freedman, R. (1981a).J. Stud. Alcohol 42,908-917. Sorenson, S., Dunwiddie, T., McLearn, G., Freedman, R., and Hoffer, B. (1981b). Pharmacol., Biochem. Behav. 14, 227-234. Spuhler, K., Hoffer, B., Weiner, N., and Palmer, M. (1982). Pharmacol., Biochem. Behav. 17, 569-578. Standen, N. B. (1981). Nature (London) 293, 158-159. Strahlendorf, H. K., and Strahlendorf, J. C. (1983). Neurobehav. Toxicol. Teratol. 5,221-224. Suzdak, P. D., Schwartz, R. D., Skolnick, P., and Paul, S. M. (1986a). Proc. Nutl. Acad. Sci. U.S.A. 83, 4071-4075. Suzdak, P. D., Glowa, J. R., Crawley, J. N., Schwartz, R. D., Skolnick, P., and Paul, S. M. (1986b). Science 234, 1243-1247. Suzdak, P. D., Schwartz, R. D., Skolnick, P., and Paul, S. M. (1988). Bruin Res. 444,340345.

348

FORREST F. WEIGHT

Svensson, T. H., and Engberg, G. (1980). I n “Biological Effects of Alcohol: Advances in Experimental Medicine and Biology” (H. Begleiter, ed.), pp. 535-550. Plenum, New York. Tahakoff, B., and Hoffman, P. L. (1991). In “Clinical Textbook of Addictive Disorders” (R. Frances and S. Miller, eds.). Guilford, New York (in press). Ticku, M. K., and Kuikarni, S. K. (1988). Pharmacol., Biochem. Behau. 30, 501-510. Tillotson, D. (1979). Proc. N d l . Acad. Sci. U.S.A. 76, 1497-1500. Treistman, S. N., and Grant, A. J. (1990).Altohol.: Clzn. Exp. Res. 14, 595-599. Treistman, S. N., and Wilson, A. (1987a).J. Neurosci. 7, 3207-3214. Treistman, S. N.. and Wilson, A. (1987b). Proc. Natl. Acad. Sci. U.S.A. 84, 9299-9303. Tsien, R. W. (1987). In “Neuromodulation: T h e Biochemical Control of Neuronal Excitability“ (L. K. Kaczmarek and I. B. Levitan, eds.), pp. 206-242. Oxford Univ. Press, New York. Turner, G. L., and Oldfield, E. (1979). h’dure (Loladon) 277, 669-670. Twombly, D. A., Herman, M. D., Kye. C. H., and Narahashi, T. (1990).J. P h a m c o l . Exp. T h . 454, 1029-1037. Verbanck, P., Seutin, V.,Dresse, A,, Scuvee, J., Massotte, L., Giesbers, I., and Kornreich, C. (1990). Alcohol.: Clin. Exp. Res. 14, 728-735. Wafford, K. A,, Burnett, D. M., Dunwiddie, T. V., and Harris, K. A. (1990). Science 249, 291-293. Wayner, M. J., Gawronski, D., and Roubie, C. (1971). Physiol. Behuu. 6, 747-749. Weight, F. F., and McCort, S. M.(I981j. In “Ion Selective Microelectrodes and Their Use in Excitable Tissues” (E. Sykova, P. Hnik, and L. Vyklicky, eds.), pp. 351-354. Plenum, New York. Weight, F. F., and Salmoiraghi, G. C. (1968).Adu. Phurmacol. 6, 395-413. Weight, F. F., Lovinger, D. M., White, G., and Peoples, R. W. (1991). Ann. N.Y. Acad. Scz. 625.97- 107. White, G. (1990). J . Neurophysiol. 64, 57-63. White, G., Lovinger, D. M., and Weight, F. F. (1989). Proc. Null. Acad. Sci. U.S.A. 86,68026806. White, G., Lovinger, D. M., and Weight, F. F. (1990). Bruin Rps. 507, 332-336. White, G., Lovinger, D. M., and Weight, F. F. (1992). (submitted for publication). Whiting, P., McKernan, R. M., and Iversen, L. L. (1990). Proc. Natl. Acad. Sci. U.S.A. 87, 9966-9970. Wong, E. H. F., and Kemp, J. A. (1991). Annu. Rev. Phurmacol. Toxicol. 31, 401-425. Woods, J. H.. K w k , W., and Ornstein, P. (1987). In “Excitatory Amino Acid Transmission” (T. P. Hicks, D. Lodge, and H. McLennan, eds.), pp. 205-212. Liss, New York. Woodward, J. J., and Gonzales, R. A. (1990). J. Neurochtm. 54, 712-715. Wozniak, K. M., Pert, A,, and Linnoila, M. (1990). Eur. J. Phannacol. 187, 287-289. Zbicz, K. L., and Weight, F. F. (1982). SOC.Neurosci. Abstr. 8, 91 1.

INDEX

A A-current, alcohol and, 307 Acetylcholine alcohol and, 308, 333 clonidine and, 59, 96 antiwithdrawal effects, 75, 77 neurotransmitters, 68-72 pharmacologic actions, 87-90, 92 GABA, receptors and, 206 leech nervous system development and, 121, 162, 177 olfaction and, 38 Acetylcholinesterase clonidine and, 88-89, 92 leech nervous system development and, 163 Action potential generation, alcohol and, 303-305 Activation alcohol and, 310, 330,332, 335, 339 GABA, receptors and inhibition, 203 synchronization, 264, 277 tetanization, 242, 248, 257, 263 olfaction and, 23, 31, 33 Adaptation, olfaction and, 32-33 Adenylate cyclase GABA, receptors and, 252-253 olfaction and biochemistry of transduction, 8-9, 11-13 receptor cells, 16 transfer of information, 25, 30-3 1, 34 Adrenergic neurons, clonidine and, 96 antiwithdrawal effects, 81-83 neurotransmitters, 64, 66-67 pharmacologic actions, 89, 91-92, 95 a-Adrenergic receptors, clonidine and, 56, 77, 85, 91 349

Afterhyperpolarization, alcohol and, 304305 Alcohol, 289-290, 336, 341-342 ion channels, 337-339 behavioral effects, 339-34 1 electrical excitability, 303-31 1 excitatory transmission, 3 I 1-325 inhibitory transmission, 325-332 synaptic transmission, 332-335 lipid theory, 290-292 neuronal firing, 292, 301-303 cerebellum, 292-295 hippocampus, 295-297 locus coeruleus, 297-298 raphe nucleus, 300-301 substantia nigra, 298-299 ventral tegmental area, 298-300 Alcohol withdrawal, clonidine and, 84-85 Alternating SCP (AS), leech nervous system development and, 165, 181-183 Alzheimer’s disease, clonidine and, 92-95, 99 Amiloride, olfaction and, 15-16 Amino acids alcohol and, 312, 332, 335, 337 clonidine and, 72, 96 olfaction and, 10, 17 7-Aminobutyric acid (GABA) alcohol and inhibitory transmission, 325-326, 328-332 ion channels, 337-338, 341 olfaction and, 6, 24-25 4-Aminopyridine, GABA, receptors and control of excitability, 213-218 inhibition, 199 synchronization, 268-269, 272-277 tetanization, 225, 228 AMPA alcohol and, 312 tetanization and, 234, 238, 261

350

INDEX

.-ini)gtlala. GABA, receptors and, 213 Anesthesia. alcohol and, 336, 339-34 1 excitatorv transmission, 3 11, 324-325 lipid theory, 290-292 neuronal firing. 293, 296-300. 302303 Annulus ereriot- neurons, leech nervous Eysteni development and, 173- 175 Aiitrroanterior nerve, leech nervous system development and. 169- 170 .411tilx)dies clonitlinr and, 62 GAB.&,, receptors and, 197-198, 251 leech nervous system development and, 119, 121, 15.5. 163, 165, 181 olfaction and. 4, 17-18 Antigens, olfaction and, 5 Antihypertensive effects, clonidine and, 56-57, 60. 97-98 antiwitlidraid effects, 82 iieurOti'ansniitters, 63-72 pharniacologic actions, 83 .\ntiwithdrawal effects. clonidine and, 7345, 97 .4p/ssio

alcohol and, 306-307, 309 GABA, receptors and, 267 APV alcohol and, 339 GABA, receptors and. 2 14. 220, 223, 260, 262, 270 AI'P iilc0110l and. 331, 335, 337-358. 34 I GABA., receptors and, 244-247, 250, 2.52, 2.54 olfaction and. X ATP-T-S.(;AHA, receptors and. 245, 247 Atropine alcohol and. 308 clonidine and, 80-81, 86, 88-91 .&xons alcohol and, 298. :305-306 G.ZR:\,, receptors arid, 19i-199, 207, 212 leech nervous s) stem development and. 117-118, 122 morphological differeiitiation, 1681i t i

neuorogenesis. 177, 1 i9- 180

neurochemical differentiation, 163, 16.5 olfaction and, 2, 4-7

B Basal bodies, olfaction and, 3 Basal cells. olfaction and, 2, 4 Basket cells, GABA,&receptors and, 197, 199-201, 235-236 Behavior alcohol and, 335, 339-341 leech nervous systeni development and, 125-126, 134-137, 187 Benzodiazepiane alcohol and, 328, 332, 337, 340 <;ABA, receptors and, 251-252 Bicuculline alcohol and, 328 GARA, receptors and, 206, 226 control of excitability, 207-208, 21 1, 216-217 synchronization, 268-269 Biogenic aniines clonidine and, 63-66, 83 leech nervous system development and, 121

Blast cells, leech nervous system development and, 131, 184, 186 cell lineage. 139, 141, 143, 146-148, 150 gangliogenesis, 157, 162 Blastomeres, leech nervous system development and, 129-130, 146, 185-186 Blood pressure, see nlro Hypertension; Hypotension clonidine and. 79-80, 98-99 Bromo-cuclic AMP (;ABA,, receptors and, 255-254 olfaction and, 1.5 Bursting interneurons, GABA, receptors and, 216-218 Bursts, suppression of, GABA,+ receptors and, 266 Btttanol behavior effects of, 338 cellular mechanisms and, 307, 335 excitatory transmission and, 31.5, 317, 323

INDEX C

CAI region alcohol and cellular mechanisms, 304-305 excitatory transmission, 312, 32 1 inhibitory transmission, 328 neuronal firing, 295-296 GABA, receptors and control of excitability, 209-210, 214, 218 inhibition, 197-200, 203, 206 intracellular regulation, 242, 248, 250, 254, 256 LTP, 232,234-236,238-241 synchronization, 267, 269-277 tetanization, 222-223, 225-230, 258260,262-263 CA3 region alcohol and cellular mechanisms, 304-305, 312, 328 neuronal firing, 296-297 GABA, receptors and control of excitability, 208-214, 218 inhibition, 199-201, 207 LTP, 232, 234-235,238-239, 241 synchronization, 264, 266-271, 273277 tetanization, 219, 222, 226, 228, 259 Calcineurin, GABA, receptors and, 246247, 259,261 Calcitonin, olfaction and, 7 Calcium alcohol and, 336 cellular mechanisms, 304, 306, 30831 I excitatory transmission, 3 15 inhibitory transmission, 331-332 clonidine and, 75 GABA, receptors and, 196, 205 intracellular regulation, 244-25 1, 254,257 synchronization, 267--269 tetanization, 228-229, 258, 260-263 leech nervous system development and, 168 olfaction and, 8, 11, 14-15, 32-33 Calmodulin GABA, receptors and

35 1

intracellular regulation, 242, 246249, 251 tetanization, 259 olfaction and, 11 Calmodulin-dependent kinase, GABA, receptors and, 249-25 1, 26 1 Calpain, GABA, receptors and, 261 Candidate receptor molecules, olfaction and, 18-19 Cardiovascular actions, clonidine and, 98 antiwithdrawal effects, 74-77, 80, 82 neurotransmitters, 64-69, 71-72 Catecholamines, clonidine and, 60, 74, 96-97 neurotransmitters, 64, 66, 72 pharmacologic actions, 85, 90-94 Caudal alternating SCP (CAS), leech nervous system development and, 165, 181-183 cDNA, olfaction and, 39 Cell lineage, leech nervous system development and segmental neurons, 139- 146 supraesophageal ganglion, 146- 147 tracing, 137-139 transfating, 147-151 Cellular mechanisms, alcohol and electrical excitability, 303-3 1 1 excitatory transmission, 3 11-325 inhibitory transmission, 325-332 synaptic transmission, 332-335 Central nervous system alcohol and, 339, 341 cellular mechanisms, 31 1-313, 315, 333 neuronal firing, 297, 301-302 clonidine and, 63, 78, 84, 96 GABA, receptors and, 195-196, 207 synchronization, 264 tetanization, 241-242, 247, 250, 256, 259 leech nervous system development and, 115, 117-120, 185 cell lineage, 139, 146-147 differentiation, 163, 168 gangliogenesis, 155-157 olfaction and, 2 Cerebellum, alcohol and, 292-295, 329 Cerebral cortex, alcohol and, 301, 326

352

INDEX

Cerebrospinal fluid. clonidine and, 83, 94, 96 Chloral hydrate, alcohol and, 297-300, 340 Chloride alcohol and, 328-331 GABA, receptors and, 196, 198-199 control of excitability, 208, 2 15-2 16 intracellular regulation, 242, 248, 252-2.53, 2.57 LTP, 238-239 physiology, 202, 206-207 olfaction and, 12. 14-15. 38 Chlorpromazine. clonidine and, 56 Choline acetyltransferase clonidine and, 77, 92 leech nervous system development and, 119. 184 Cholinergic junctions, alcohol and, 333 Cholinergic neurons clonidine and, 96-99 antiwithdrawal effects. 74-55, 80-84 neurotransmitters, 68-72 pharmacologic actions. 86-87, 92-93 leech nervous system development and, 119. 184 Cholinesterase. clonidine and, 69, 91, 93 Cholinesterase inhibitor toxicity clonidine and, 87, 91 Chromatograph) leech nervous system development and, 121 olfaction and. 37 Cilia, olfaction and, 3-4, I5 biochemistry of transdurtion, 8-9, 1 112 receptor patterns of response. 18, 20 transfer of information, 32, 34 Cleavage, leech nervous system development and, 184-186 cell lineage. 138. 147 morphology, 129- 131 Clones alcohol and, 338 leech nervous system development and. 146. 1.50, 157, 184, 186 olfaction and, 10, 39. 39-40 Clonidine antiwithdrawal effects, 84-85 clinical indication, 57-58

development of, 56-57 future directions, 98-99 growh hormone secretion, 85-86 inhibition of cholinesterase inhibitor toxicity, 87-92 learning and memory, 92-95 neuromodulation, 97-98 neurotransmitters, 72 acetylcholine, 68-72 biogenic amines, 63-66 opiates, 66-68 opiate withdrawal, 73-76 spinal cord model, 76-84 pharmacologic actions, 95-97 receptor specificity central sites, 60-63 peripheral sites, 59 Clonidine-displacing substance, 61-62 CNQX, GABA, receptors and, 214, 270 Connective nerves, leech nervous system development and, 171, 174-175, 180, 183, 186 Connectivity, GABA, receptors and, 228, 266-267 Convergence, olfaction and, 34-35 Cyclic AMP clonidine and, 97 GABA, receptors and, 250-254, 259 olfaction and, 7-11, 15-16, 33-34 Cyclic nucleotides, olfaction and, 9, 15-16 Cvtoplasnt leech nervous system development and, 129 olfaction and, 4 Cytoskeleton, olfaction and, 3

D Dantrolene, alcohol and, 31 1 DAKPP, GABA, receptors and, 259, 26 1 Delayed rectifier current, alcohol and, 306-307 Dendrites GABA,, receptors and control of excitability, 209, 2 16 inhibition, 197-199, 202, 205, 207 intracellular regulation, 242, 246 LTP, 232-234, 237-238, 240-24 I

INDEX

synchronization, 266, 274 tetanization, 223, 263, 267 olfaction and, 3, 6-7, 9, 15 Dendrodendritic synapses, olfaction and, 6- 7 Dentate gyrus, GABA, receptors and, 200, 218,238, 259,270 2-Deoxyglucose, olfaction and, 23-24, 29 Dephosphorylation, GABA, receptors and intracellular regulation, 241-247, 249, 252 tetanization, 258-259, 261 Depolarization alcohol and, 312 GABA, receptors and control of excitability, 21 1-212, 214218 inhibition, 199, 202-203, 206 LTP, 231, 233, 235,237-238, 241 synchronization, 266-268, 270-273, 275,277 tetanization, 220, 222-223, 225-230 leech nervous system development and, 167-1 68 olfaction and, 21, 33, 38 Desensitization, GABA, receptors and, 253 Diacylglycerol, GABA, receptors and, 254, 256 Differentiation GABA, receptors and, 230 leech nervous system development and, 180, 184 electrophysiology, 167-168 morphology, 168- 176 neurochemistry, 162-167 olfaction and, 4-5 Direct gating, olfaction and, 12 Disinhibition, GABA, receptors and control of excitability, 208-2 13 LTP, 233,240-241 physiology, 205 synchronization, 266-268, 274, 276 tetanization, 259-260 DNA, leech nervous system development and, 156 DNase, leech nervous system development and, 148 Dopamine

353

alcohol and, 298-300, 326,340 clonidine and, 56 leech nervous system development and, 120, 143, 158, 163, 165, 185 Dorsal root ganglion (DRG) neurons, alcohol and, 337 cellular mechanisms, 305, 309-310 excitatory transmission, 3 13, 323 inhibitory transmission, 330 Dorsoposterior nerve, leech nervous system development and, 115, 169-170, 172

E Effector neurons, leech nervous system development and, 171-172, 175-176 EGTA, alcohol and, 309-310 Electrical excitability, alcohol and, 303311 Electron microscopy, olfaction and, 3 Electrophysiological differentiation, leech nervous system development and, 157-158 Embryo, leech nervous system development and, 113, 185-187 behavior, 134, 136-137 cell lineage, 138-139, 147-148, 150151 differentiation, 162, 167-173 gangliogenesis, 154-156, 158-159, 161-162 morphology, 127, 129, 131-132, 134 neuorogenesis, 177, 179- 181 Embryogenesis, leech nervous system development and, 111, 138, 175, 183 behavior, 134, 137 morphology, 130, 132 P-Endorphin, clonidine and, 67-68, 72 Entrainment, inhibitory, GABA, receptors and, 267-268 Enzymes clonidine and, 64, 77, 87-88, 92, 95 GABA, receptors and, 196, 206, 241, 245, 252, 261 olfaction and biochemistry of transduction, 9, 1112 perireceptor events, 38-39

354

INDEX

receptor patterns of response, 17 transfer of information, 30, 32-33 Epileps! alcohol and. 3 13 (;ABA, receptor-s and control o f excitability, 208, 213 synchronization. 264, 266, 269, 271, “3-274 tetanization, 220, 259 Epinephrine. clonitfine and. 61, 67 Epitheli um <;;\BA,, receptors and, 2 19 leech nervous system development and, 1 3 1 . 145, 150. I59 oltaction anti. 2. 4-6 lhchcniistry o f transduction, 8-9, 12 perirereptor events, 3;-38, 40 receptor cells, I4 receptor patterns of response, 18, 20-22 Iranster of information, 29 E-S potentiation. GABA,.\ receptors and, 2.3-24 1 Ethanol cellular mechanisnrs and, 303-31 1 rxcitatory transmission and, 31 1-313, 31.5. 317. 319. 321-325 inhibitor) transmission and, 325-326. 328-332 ion channels a i d . 336-342 neuronal firing ancl, 292-303 s! naptic transmission and, 333-335 Excitation alcohol and, 33.5-337, 339, 34 1 cellular mechanisms, 303, 308, 31 1 excitator) transmission, 3 1 1-325 neuronal firing. 293, 296-301 clonidine ancl antirvithdrawl effects, 74-77. 81-82, 84

Iretlrotraiisinitters, 68, 72 pharnracologic actions, 86 (;ABri,\ receptoi-s and, IY6-197 anatomy. 198. 200-201 control, 207-208 disinhibition. 208-2 13 inhibitory circuit, 213-218 I-TP, 23 1 , 233, 235-238, 24 1 physiology, 202-205. 207 synchronization, 266-277

tetanization, 219, 226-228, 257, 259263 leech nervous system development and, 118-119, 121, 126, 168, 177 olfaction and, 6, 21-23, 34 Excitatory postsynaptic potentials (EPSPs) alcohol and, 304, 312, 319, 321, 325 GABA, receptors and control, 209-212, 214, 216, 218 inhibition, 199-200, 202-205, 207 LTP, 233-241 synchronization, 274-275 tetanization, 219-220, 222, 225, 228

F Fatty acids, olfaction and, 13 Feedback, GABA, receptors and, 197, 199, 201, 218, 263, 266 b’eetlforward, GABA,, receptors and, 197, 218 inhibition, 199-201 tetanization. 233, 236, 239, 241 Fertilization. leech nervous system development and, 114, 127 Fluorescein-dextran tracer, leech nervous system development and, 138-139, 150

Fluorescence, leech nervous system development and, 139-120, 138, 163, 165, 167 Flurazepani, alcohol and, 328, 330-332 FMRFamide, leech nervous system development and, 121, 165 Formaldehyde, leech nervous system development and, 119, 121 Forskolin GABA,, receptors and, 253 olfaction and, 8, 16 Freeze-fracture studies, olfaction and, 4 Furosemide, olfaction and, 15

G G proteins, SPP Guanine nucleotide binding proteins GABA (7-Aminobutyric acid) alcohol and

INDEX

inhibitory transmission, 325-326, 328-332 ion channels, 337-338, 341 olfaction and, 6, 24-25 GABA, receptors, 195-1 97 function, control of excitability and, 207-208 disinhibition, 208-213 synchronization, 213-218 function, synchronization and, 263266, 273-278 disinhibition, 266-268 inhibitory circuit, 268--270 tetanization, 271-273 function, tetanization and, 257-263 features, 218-230 intracellular regulation, 24 1-257 LTP, 231-241 GABAergic inhibition, anatomy and circuitry, 199-201 distribution, 198- 199 interneurons, 197-198 GABAergic inhibition, physiology and, 202-207 GABAergic neurons, leech nervous system development and, 119, 165 Gangliogenesis, leech nervous system development and, 154-163 Ganglion alcohol and, 304-306, 309, 333, 335 leech nervous system development and, 115-122, 126, 177, 179-180, 183 cell lineage, 139, 141, 146-147 gangliogenesis, 154- 162 morphological differentiation, 168173, 175 morphology, 132, 134 neurochemical differentiation, 162163, 165, 167 GDP, olfaction and, 8 Geneology, leech nervous system development and, 139-146, 183-185 Genetic factors, alcohol and, 294-295 Glial cells GABA, receptors and, 196, 246 leech nervous system development and, 110, 116, 167, 184 cell lineage, 141, 143 gangliogenesis, 157-159 olfaction and, 11

355

Glial fibrillary acidic protein (GFAP), olfaction and, 4 Glomerulus, olfaction and, 5-7, 23-24 Glossiphoniid leeches, nervous system development of behavior, 137 differentiation, 167-168 gangliogenesis, 154 morphology, 129-130, 132, 134 myogenesis, 154 neuorogenesis, 181 Glutamate alcohol and, 311-325,339 clonidine and, 76 leech nervous system development and, 119 Glutamic acid decarboxylase (GAD), GABA, receptors and, 198, 206 Glycine, alcohol and, 312, 319, 326, 329, 337, 341 Glycoprotein, olfaction and, 18 Glyoxylic acid, leech nervous system development and, 119-120, 163, 165 Growth-associated protein, olfaction and, 5 Growth hormone secretion, clonidine and, 84-85 GTP, olfaction and, 8-10, 12 GTPase, olfaction and, 18 Guanine nucleotide binding proteins (G proteins) alcohol and, 335 GABA, receptors and, 199, 203 olfaction and, 8-10, 12, 15, 32, 34

H Haementeria, leech nervous system development and, 113-1 14, 117, 120 behavior, 134, 136-137 differentiation, 162-163, 165, 173 morphology, 127, 129, 132 Haemopis, leech nervous system development and, 155-156 Halothane, neuronal firing and, 293, 296-297,299 Heart accessory neurons, leech nervous system development and, 173-174 Helix, alcohol and, 309

INDEX

Helobdella, leech nervous systern development and, 113. 120 cell lineage, 148, I50 differentiation. 165 morphology, 127, 129, 132 neuorogenesis, 183 Hemicholiniuni-3. clonidine and, 71, 75, 80-8 1 Hemisegments, leech nervous system development and, 143, 146, 151 Hippocampus, SPY a1.w CAI region; CX3 region alcohol and. 337 cellular mechanisms, 304-305, 308, 3 10 excitatory transmission, 3 12-3 13, 31.5, 319, 321, 323 inhibitor) transmission, 328. 330 neuronal firing, 295-297 clonidine and, 99 GABA, receptors and, 197 control of excitability, 208-209. 2 12217 inhibition, 197-201. 203. 205-207 intracellular regulation, 242, 244, 246, 249, 253-2.54, 256 i n , 231-234, 238 synchronization, 266, 268-27 I , 274, 276-277 tetanization, 218-220, 222. 227, 258, 26 1-263 I-lirudo, leech nervous system development and, 1 1 1 , 114, 117, 121 differentiation, 162-163. 172-173, I75 gangliogenesis, 156 morphology, 132, 134 myogenesis. 1.53 neuorogenesis, 177. 179, 183 Histamine. olfaction and, 38 Homology, leech nervous system development and, 116, 119, 127-134 behavior, 134 cell lineage, 146 differentiation, 168- 176 gangiiogenesis. 155, 157 netlowgenesis. 177 I formones clonidine and, 73, 86, 92 olfaction and, 7. 16, 28, 30 Horseradish peroxidase, leech nervous

system development and. 138, 150, 168

Hydrolysis, GABA, receptors and, 245, 250 Hydrophobicity, alcohol and, 317, 338, 342 6-Hydroxydopamine, alcohol and, 293, 298 5-Hydroxytryptarriine channels, alcohol and, 335-338, 340 Hyperpolarization GABA,* receptors and, 196 control of excitability, 21 I , 214-215, 218 inhibition, 199, 201-203 LTP, 233, 235-236, 238 synchronization, 266-268, 271, 277 tetanization, 220, 223, 225, 228 olfaction and, 2 1 Hypertension, clonidine and, 98 antiwithdrawal effects, 75 neurotransmitters, 63, 67-71 receptor specificity, 61 Hypotension, clonidine and, 57, 97 neurotransmitters, 64, 72 receptor specificity, 59, 61 Hypothalamus alcohol and, 301-302 clonidine and, 60-61, 69, 7 1, 86

I Iniidazole, clonidine and. 61-62, 96 lniniunoreactivity leech nervous system development and, 121, 163, 165, 181 olfaction and, 37 Inhibition alcohol and, 335, 337-339, 341 cellular mechanisms, 305, 309-3 11 excitatory transmission, 3 13, 3 15, 317, 319, 321-325 inhibitory transmission, 325-332 lipid theory, 291 neuronal firing, 293-296, 298, 300302 clonidine and, 56-57, 96-97, 99 antiwithdrawal effects, 74-75, 77, 8084

INDEX

neurotransmitters, 64-65, 67, 69, 7172 receptor specificity, 59-62 GABA, receptors and, 195-197 anatomy, 197-201 control of excitability, 207-216 intracellular regulation, 242, 245246,251, 253 LTP, 231,233-241 physiology, 202-207 synchronization, 264, 266-273, 275278 tetanization, 219-220, 223, 228, 259260, 263 leech nervous system development and, 119, 121, 165, 168, 172 olfaction and, 8-1 1, 22, 30, 34 Inhibitory entrainment, GABA, receptors and, 267-268 Inhibitory postsynaptic currents (IPSCs) alcohol and, 328 GABA, receptors and, 205, 220, 277278 Inhibitory postsynaptic potentials (IPSPs) alcohol and, 304,328, 341 GABA, receptors and, 196 anatomy, 198-201 control of excitability, 209-21 1, 214218 intracellular regulation, 249 LTP, 234-235,237,239-240 physiology, 202-207 synchronization, 267-273, 275-278 tetanization, 219-220, 222, 225-228, 230, 260, 263 Inositol triphosphate, GABA, receptors and, 248, 250 Insulin, olfaction and, 7 Intermediate filament protein, olfaction and, 4 Interneurons alcohol and, 302 GABA, receptors and, 197 control of excitability, 2 15-2 18 inhibition, 197-201, 205-206 LTP, 233,235-237 synchronization, 268-273, 275-277 tetanization, 222, 225-229, 260 leech nervous system development and, 172-173, 180

357

Intoxication cellular mechanisms and, 315, 317, 324 ion channels and, 339-341 lipid theory of alcohol and, 291 neuronal firing and, 297 Intracellular regulation, GABA, receptors and, 241-259 Inward rectifier current, alcohol and, 308 Ion channels GABA, receptors and, 241, 248, 250, 256 olfaction and, 14-16, 33 Ion gates, olfaction and, 12, 14-15, 27, 32, 34 Isoenzymes, olfaction and, 11, 39 Isopentanol, synaptic transmission and, 315, 327, 335, 338

K Kainate, alcohol and excitatory transmission, 312-313, 319, 323,325 ion channels, 337,339-340 Ketamine, alcohol and, 319 Kinship groups, leech nervous system development and, 141, 143, 146, 148, 184-185 Korsakoff’s disease, clonidine and, 9495

L Lacunosum/moleculare interneurons, GABA, receptors and, 197-198, 200-201 Learning alcohol and, 295, 313 clonidine and, 92-94, 99 GABA, receptors and, 219, 231-232 olfaction and, 24-25 Lectins, olfaction and, 6, 17 Leech nervous system development, 109110, 183-187 behavior, 124-126, 134-137 cell lineage segmental neurons, 139- 146 supraesophageal ganglion, 146-147

358

INDEX

tracing, 137-139 transtating, 147- 1 .il differentiation electrophysiolo~ical, 167- 168 morphological. 168- 176 neurochernical, 162- 167 gangliogenesis. 154- 162 gross anatom); 1 12- 1 15 history, 1 10- 1 1 1 identified cells. I 17- 1 I9 identified circuits, 127 morphology., 127- 134 ni\-ogenesis, 1.5 1- I54 neuorogenesis. 176- 183 neurotransmitters. 119- 121 sensory and motor fields, 122- 123, 125 taxonomy, 1 1 1, 113 ventral nerve cord. I 15- 1 17 Ligands alcohol and, 312. 332 clonidine and, 6 I GABA,., receptors and, 241, 248, 253, 256 olfaction and, 8, 10, 12, 3 1 Lipid theory of alcohol, 280-292, 307, 336, 338, 34 1 Lipids clonidine and, 91 olfaction and. 8, 10. 12-13, 29 Liposomes, olfaction and. 12 Lobster, olfaction and. 19, 38-39 Locomotion, leech nei-vous system development and, 125, 137 Locus coeruleus alcohol and, 297-298, 3 1 3 clonidine and. 74 Long-lasting depolarizing potential (LLD), G A B A 4 receptors and, 215, 217. 269, “7 Long-sleep mice. alcohol and, 295-296, 332, 337 Long-term potentiation (LTP) alcohol and, 313 GABA receptors and, 2 18-2 19, 23 124 1, 25Yr 26 1-263 Longitudinal fibers, leech nervous system developnient and, 1.53, 171172

,

M M-current, alcohol and, 307-308, 336 Macromeres, leech nervous system development and, 130, 132 Magnesium alcohol and, 312, 319 GABA, receptors and, 233, 244, 247, 254 Mean arterial pressure, clonidine and, 78 Mechanosensory neurons, leech nervous system development and, 168- 171, 173, 177 Medial septa1 nucleus, alcohol and, 328 Medioanterior (MA) nerve, leech nervous system development and, 115, 169171 Medulla, clonidine and, 60-61, 69, 82 hiembrane currents, olfaction and, 16 Membrane potential, alcohol and, 312 Membrane potential oscillations (MPOs), GABA, receptors and, 228-230, 273 Membrane properties, alcohol and, 303305,308-309 Memory alcohol and, 295 clonidine and, 92-94, 99 GABA,%receptors and, 231, 259 Metacerebral cells (MCC), alcohol and, 307 Methanol, excitatory transmission and, 315. 323,338 Micromeres, leech nervous system development and. 130-131, 147 Microtubule-associated proteins, olfaction and. 4 Migration, leech nervous system development and, 134, 139, 156-158, 186 Mitosis, leech nervous system development and, 183, 186 hlonoclonal antibodies, olfaction and, 6, 18

Morphine alcohol and, 340 clonidine and, 67, 73-83 Morphogenesis, leech nervous system development and, 110, 131, 151, 173 Morpholog clonidine and, 94

INDEX

GABA, receptors and, 212, 260-261 olfaction and, 2, 7, 37 Motor neurons, leech nervous system development and, 172-175, 184 mRNA GABA, receptors and, 256 olfaction and, 9-10, 14 Muscarinic receptor alcohol and, 308 clonidine and, 97 antiwithdrawal effects, 75, 77, 80-81 neurotransmitters, 69-72 pharmacologic actions, 87-89 Muscimol, GABA, receptors and, 253 Muscle fibers, leech nervous system development and, 171-172 Myoblasts, leech nervous system development and, 151, 153-154 Myogenesis, leech nervous system development and, 136, 151-154

N Naloxone, clonidine and, 67-68, 78, 808 1, 83-84 Neocortex alcohol and, 315 CABA, receptors and control of excitability, 213-214 synchronization, 268, 270, 274 tetanization, 233, 252, 256 Neostigmine, clonidine and, 81, 87 Neostriatum, alcohol and, 298 Neuorogenesis, leech nervous system development and, 147, 156, 176, 179180 Neuroblastoma cells, alcohol and, 310,333 Neurochemistry, leech nervous system development and, 158, 161-167, 182 Neuromodulators, clonidine and, 97-98 Neuronal firing, alcohol and, 292-303 Neuropeptides clonidine and, 72 GABA, receptors and, 198 leech nervous system development and, 121, 165 Neuropil, leech nervous system development and, 116, 120

359

differentiation, 168-171, 173, 175 gangliogenesis, 159 neuorogenesis, 177 Neurotoxicity alcohol and, 313 clonidine and, 66, 86 Neurotransmitters alcohol and, 333, 337, 341 cellular mechanisms, 303, 308 excitatory transmission, 312, 315 clonidine and, 56, 63, 72, 96-97, 99 acetylcholine, 68-72 antiwithdrawal effects, 74-76 biogenic amines, 63-66 opiates, 66-68 pharmacologic actions, 86, 92-93, 95 GABA, receptors and, 252 leech nervous system development and, 119-123, 125, 143, 167, 180 olfaction and, 2, 16, 28, 30 Neurpeptides, olfaction and, 7 Nicotine, alcohol and, 340-341 Nicotinic acetylcholine receptors, alcohol and, 333, 337, 341 NMDA (N-Methyl-D-aspartate) alcohol and excitatory transmission, 3 12-3 13, 315, 317, 319,321, 323-325 ion channels, 337, 339 GABA, receptors and intracellular regulation, 249 LTP, 233-234, 241 synchronization, 276 tetanization, 220, 223, 230, 260-263 Noradrenaline, olfaction and, 24-25 Noradrenergic neurons alcohol and, 293, 297 clonidine and, 96, 99 antiwithdrawal effects, 73-74, 82 neurotransmitters, 63-66 pharmacologic actions, 91, 95 Norepinephrine, clonidine and, 59-62, 74, 94, 96 Nucleus accumbens, alcohol and, 298, 340 0 1-0ctano1, cellular mechanisms and, 3 11 Odor concentration, olfaction and, 26-28, 33

360

1N DEX

Odorant-binding protein, olfaction and, 39-40 Olfaction. 1-2. 40-4 1 biochemistry of transduction, 7-8 cyclic AMP, 8- 1 1 direct gating, 12 lipid responses, 12-13 phosphoinositides, 1 1 - 12 perireceptor events, 35 control of secretion, 37-38 control of sensory cells, 38 odorant-binding protein, 39-40 olfactory mucus, 35-37 xenobiotic-metabolizing enzymes, 3839 receptor cells, 13- 16 receptor patterns of response, 16 olfactory bulb, 20-25 receptor cells, 19-20 receptor molecules, 17-19 Ytructure, 2-7 transfer of information, 25 from mucus to receptor molecule, 28-32 odor concentration, 26-28 from receptor molecule to cell, 32-35 vectorial representation. 25-26 Olfactory bulb, 6-7, 20-25 Olfactory marker protein (OMP), 5 Olfactory mucus, 35-37 Opiates, clonidine and, 98 antiwithdrawal effects, 73-84 neurotransmitters, 66-68, 72 Oriens/alveus interneurons, GABA, receptors and, 199, 201 Oxygen. leech nervous system development and, 139

P Pargyline, clonidine and, 90-91 P m compdcta neurons, alcohol and, 298299 Penicilline. (;.4BA, receptors and, 208, 21 I Pentobarbital, alcohol and, 299, 302, 31 I , 326,330-331 Pentobarbitol, GABA,, receptors and, 199, 216

Peptides, clonidine and, 73, 76, 83 Peripheral nervous system, leech nervous system development and, 139, 141, 158- I59 Peripheral targets, leech nervous system development and, 176- 180 Perireceptor events, olfaction and, 35-40 Peristalsis, leech nervous system development and, 134- 135 Pharmacology alcohol and, 291, 31 1, 336-337, 341 clonidine and, 56-58, 95-99 antiwithdrawal effects, 73-74, 79, 82, 85 growth hormone secretion, 85-86 inhibition of cholinesterase inhibitor toxicity, 87-92 learning, 92-95 neurotransmitters, 64-65 receptor specificity, 60 GABA,, receptors and control of excitability, 207-208 physiology, 203 synchronization, 266, 271 tetanization, 219, 260 leech nervous system development and, 119 Phencyclidine, alcohol and, 317 Phenotype leech nervous system development and, 184, 186-187 differentiation, 165 gangliogenesis, 158-159 neuorogenesis, 177, 182-183 olfaction and, 7 Phenylephrine, alcohol and, 301 Phorbol- 12.13-dibutyrate (PDBu), GABA, receptors and, 256-251 Phosphatase, GABA, receptors and, 241242, 245-249, 258 Phosphate, GABA, receptors and, 245 Phosphodiesterase, olfaction and, 8-9, 1 I Phosphoinositides, olfaction and, 8, 1112 Phospholipase C, olfaction and, 11 Phospholipids, GABA, receptors and, 250 Phosphorylation alcohol and, 332, 339 GABA, receptors and

INDEX

intracellular regulation, 24 1-247, 249,251-254,256 tetanization, 230, 258-259, 262 olfaction and, 9, 16 Physostigmine, clonidine and, 99 neurotransmitters, 69-72 pharrnacologic actions, 87, 91, 93 Picrotoxin, GABA, receptors and, 196 control of excitability, 207-209, 21 I , 21 7-2 18 inhibition, 199, 206 synchronization, 268-269, 273-274 tetanization, 219, 249, 262 Pineal gland, alcohol and, 307 Platelets, clonidine and, 97 Population EPSPs, alcohol and, 312, 321, 323 Posteroposterior (PP) nerves, leech nervous system development and, 115, 169-171 Postsynaptic potentials, see Excitatory postsynaptic potentials; Inhibitory postsynaptic potentials Potassium alcohol and, 304, 306-309, 31 1 GABA, receptors and control of excitability, 211, 215, 218 inhibition, 198-199, 202-203, 205 synchronization, 267 tetanization, 223, 228 olfaction and, 12, 14, 35 I-Propanol, alcohol and, 335, 338 Propranolol, alcohol and, 293 Proteases, GABA, receptors and, 245, 26 1 Protein alcohol and, 291-292, 338, 342 GABA, receptors and inhibition, 198 intracellular regulation, 241-242, 246-253 tetanization, 263 olfaction and, 4-5, 29, 40 biochemistry of transduction, 8- 10, 12-13 receptor patterns of response, 17-18 Protein kinase, GABA, receptors and intracellular regulation, 241, 245, 247, 250-257 tetanization, 259, 261

36 1

Protein kinase A, GABA, receptors and, 250-254,258-259 Protein kinase C, GABA, receptors and, 250-251,254,256-258,262 Proteolysis, GABA, receptors and, 245, 26 1 Purkinje cells, alcohol and, 292-295, 326, 329 Pyramidal cells alcohol and cellular mechanisms, 304-305, 308, 312,321 neuronal firing, 295-297 GABA, receptors and control of excitability, 208-21 1, 214215 inhibition, 197-201, 207 intracellular regulation, 244, 248, 250,253,257 LTP, 232-236,238,241 synchronization, 266-27 1, 275-276 tetanization, 219, 223, 225, 228, 258259,262-263

Q Quisqualate, alcohol and excitatory transmission, 312-313, 319, 321,323, 325 ion channels, 337,339-340

R Raphe nucleus, alcohol and, 300-301 Receptor cells, olfaction and, 13-16 Receptor-mediated synaptic excitation, alcohol and, 321,323 Receptors alcohol and, 339-340 clonidine and, 56, 96-98 antiwithdrawal effects, 75-77, 80 neurotransmitters, 69-72 pharmacologic actions, 87-89, 91 specificity, 59-63 GABA,, see GABA, receptors olfaction and biochemistry of transduction, 10 olfactory bulb, 20-25

INDEX

piartrrns of response. 16-20 perireceptor- events. 36. 40 transfer of information, 25-35 Krctifiei-current. alcohol and. 306-308 Kepolariration. alcohol and, 306 Keproductive ducts. leech nervous sFsteni development and. 177, 179-180 Kespiratc~rycycle. olfaction and. 20-2 1 Ketzius cells, leech nersous system developnient and. 119, 126. 163. 165, 17:i- 177 Khodamiiie, Ire
S

Second messengers alcohol and. 33.5 clonidine and, 62 GAB;\, receptors and intracellular regulation, 24 1. 248, 253 tetanization, 2.37-258, 261-262 olfaction and, 30. 92-34 Secretion, olfaction anti, 3 - 3 9 Segniental neurons. leech nerwms system development and cell lineage. 139-146 differentiation, 168- 169. 172-173. 17~5-176 neuorogenesis. 177. 180 Segmentation, leech nervous system tfeselopmenr and cell lineage, 143, 147, 150 gangliogenesis. 155- 159. 162 morphology. 132 i n yogenesis. 1.5 1 1.54 Sensitivity GABA receptors and intracellular regulation. 235-236

,

synchronization, 276 tetanization, 222-230 olfaction and, 32 Sensory cells, olfaction and, 38 Sensory neurons, alcohol and, 306 Septa1 area, alcohol and, 301 Sequences alcohol and, 338 GAB24, receptors and, 199, 209, 219, 22.3, 250, 252 leech nervous system development and, 131,184

olfaction and, 40 Serotonin alcohol and, 300, 326, 333 clonidine and, 66, 86 leech nervous system development and, 119-120, 126, 180, 185 behavior, I36 cell lineage. 1.13 differentiation, 162-163, 165 gangliogenesis, 157, 159, 161 neuorogenesis. 180 Short-sleep mice, alcohol and. 295-296, 337 Signal/noise, olfaction and, 37-38 Signal transduction, GABA,+ receptors and, 242, 250, 254 Small cardiac peptide (SCP),leech nervous system development and, 121, 165, 181,183 Sodium (;.4BA:, receptors and, 196, 228, 244, 267 olfaction and. 12, 14-15, 35 Sodium chaniiels, alcohol and, 305-306, 309. 31 1 Soman, clonidine and, 88-92 Somarostatin, alcohol and, 308 Spinal cord alcohol and, 34 1 cellular mechanisms, 305 excitatory transmission, 3 1 1, 3 15 inhibitory transmission, 326, 328-329 neuronal firing, 301-302 clonidine and antiwithdrawal effects. 74, 76-84 neurotransmitters, 63 receptor specificity, 60-62

INDEX

363

Spontaneous EPSPs, GABA, receptors and, 222 Spontaneous IPSCs, GABA, receptors and, 205 Spontaneous IPSPs, GABA, receptors and inhibition, 205-207 synchronization, 268, 271, 276-278 tetanization, 220, 222 Spontaneously hypertensive rats, clonidine and, 63-64, 71 Steroids, olfaction and, 39 Stimulation-evoked potentials, GABA, receptors and, 218-229, 235 Strata oriens, GABA, receptors and, 197-199 Strata radiatum, GABA, receptors and, 198, 200, 219, 230, 274 Striatum, alcohol and, 315 Substance P clonidine and, 76-77, 80 olfaction and, 37-38 Substantia nigra, alcohol and, 298-299 Supraesophageal ganglion, leech nervous system development and, 146-147 Sympathoexcitation, clonidine and, 68, 74-75, 77,81-84 Sympathoinhibition, clonidine and, 74, 82 Synapses, alcohol and cellular mechanisms, 303, 332-335 excitatory transmission, 3 11-325 inhibitory transmission, 325-332 Synaptic plasticity, GABA, receptors and, 231-232, 261-262 Synaptic potentials, GABA, receptors and, 218-222 Synchronization, GABA, receptors and, 263, 273-278 disinhibition, 266-268 inhibitory circuit, 267-270 physiology, 263-266 tetanization, 232, 259, 270-272

Urethane, neuronal firing and, 293, 296, 302

T

V

Taxonomy, leech nervous system development and, 11 1, 113

Vectorial representation, olfaction and, 25-26, 29-30

Teloblasts, leech nervous system development and, 158, 181, 185 cell lineage, 139, 141, 143, 146-148 morphology, 131-132 Teloplasm, leech nervous system development and, 129-130, 186 Tetanization, GABA, receptors and, 257263 features, 2 18-230 intracellular regulation, 241-257, 271273, 276-278 LTP, 231-241 Tetrodotoxin alcohol and, 309 GABA, receptors and, 205, 218, 222, 274 olfaction and, 14 Therornyzon, leech nervous system development and, 113, 150, 165, 183 morphology, 127, 129, 131-132 Tissue interactions, leech nervous system development and, 158-162 Toxicity, clonidine and, 87-92 Transduction, olfaction and, 4, 7-13, 41 perireceptor events, 35 receptor cells, 15 transfer of information, 25, 34 Transductory apparatus, olfaction and, 3, 32,34 Transfating, leech nervous system development and, 147-151 Transmitter-gated ion channels, alcohol and excitatory transmission, 31 1-325 inhibitory transmission, 325-332 sensitivity, 339-341 synaptic transmission, 332-335

U

364

INDEX

Vemeronasal system, olfaction and, 2-3, 26 Ventral nerve cord, leech nervous system development and. 115-1 17, 139, 147, 172 Ventral tegmental area, alcoliol and, 298so0 Voltage-gated ion channels, alcohol and, 305-31 I , 336-337 W Whole cell responses, olfaction and, 1516 b.hole tnsue responses, olfaction and, 15

Withdrawal, opiate, clonidine and, 73-84, 98

X Xenobiotic-metabolizing enzymes, olfaction and, 38-39 Xenopzlr alcohol and, 332 GABA, receptors and, 256 olfaction and, 14

2 Zinc, alcohol and, 3 19

CONTENTS OF RECENT VOLUMES

Volume 24

Volume 23

Chemically Induced Ion Channels in Nerve Cell Membranes David A. Mathers and Jeffeery L. Barker Fluctuations of Na and K Currents in Excitable Membranes Berthold Neumcke Biochemical Studies of the Excitable Membrane Sodium Channel Robert L. Barchi Benzodiazepine Receptors in the Central Nervous System Phil Skolnick and Steven M . Paul Rapid Changes in Phospholipid Metabolism during Secretion and Receptor Activation F. T. Crews Glucocorticoid Effects on Central Nervous Excitability and Synaptic Transmission Eduard D. Hall Assessing the Functional Significanceof Lesion Induced Neuronal Plasticity Oswald Steward Dopamine Receptors in the Central Nervous System Ian Cresse, A. Leslie Morrow, Stuart E. Leff. David R. Siblqr and Mark W. Hamblin Functional Studies of the Central Catecholamines T. W. Robbim and B. J . Eueritt Studies of Human Growth Hormone Secretion in Sleep and Waking Wallace B. Mendelson Sleep Mechanisms: Biology and Control of REM Sleep Dennis J . McGinty and Rent R. Drucker-Colin INDEX

Antiacetylcholine Receptor Antibodies and Myasthenia Gravis B m r d W. Fulpiur Pharmacology of Barbiturates: Electrophysiological and Neurochemical Studies Max w i h and ~ (hham A. R. Johnston

,,

Immunodetection of Endorphins and Enkephalins: A Search for Reliability Akjandro William Shoemaker, Jacqueline F. McCinty, and Floyd Bloom

On the Sacred Disease: The Neurochemistry of Epilepsy 0. Carter Snead III Biochemical and ElectrophysiologicalCharacteristics of Mammalian GABA Receptors Salvatore 1.Enna and Joel P. Gallagher Synaptic Mechanisms and Circuitry Involved in Motorneuron Control during Michael H. Chase

Recent Developments in the Structure and Function of the Acetylcholine Receptor F. J. Barrantes Characterization of a,- and a,-Adrenergic Receptors David B. Bylund and David C. U'Prichard Ontogenesis of the Axolemma and Axoglial Relationships in Myelinated Fibers: Electrophysiological and Freeze-Fracture 'Orrelates of Membrane Plasticity Stephen G. Waxman,Joel A. Black, and Robert E. Foster INDEX

365

366

CYJNTENTS OF RECENT VOLLMES

Volume 25

Guanethidine-Induced Destruction of Sympathetic Neurons Eugene M . Johiaton, Jr. arid Patneb TOJMattiting Dental Sensory Receptors Margaret R . B y r s Cerebrospinal Fluid Proteins in Neurology A. Lawenthnl. R . Crols, E . De Schidter, J. Gheurexc, D . Knrcher, M . A'oppe, mnd A. TclrnieI Muscarinic Receptors in the C;entral Nervous System Mordtchai Sokol
Muscarinic Receptor Subtypes in the Central Nervous System Wayne Hoss and John Ellis Neural Plasticity and Recovery of Function after Brain Injury John F. Mardud1 From Immunoneurology to Immunopsychiatry: Neuromodulating Activity of AntiBrain Antibodies B r a n k h i D . Jariklovik

Effect of Tremorigenic Agents on the Cerebellum: A Review of Biochemical and Electrophysiologica! Data 1.' G . Long0 and M . Mmsotti INDEX

Peptides and Nociception Daniel Luttinger, Daniel E . Hernandez, Ctiarb B . Nenieroff, and ArthurJ. Pmnge. Jr.

Volume 27

Opioid Actions on Mammalian Spinal Neurons 14: Ziuglganshrt.gur

T h e Nature of the Site of General Anesthesia Keith W. Miller

Psychobiology o f Opioids Alberto Oliwtio. Claudio Custellano, and Stefan0 Pub1i.si-Allegrn

T h e Physiological Role of Adenosine in the Central Nervous System Thomm V. Dunwiddie

Hippocarnpal Damage: Effects on Dopaminergic Systems of the Basal Ganglia Robert L. I,sancson

Somatostatin, Substance P, Vasoactive Intestinal Polypeptide, and Neuropeptide Y Receptors: Critical Assessment of Biochemical Methodology and Results A nders UrLdPn, Lo u-Lou Peterson, and T ? i m Bartfni

Neurocheniical Genctics V. Csariy The Neurobiology of Some Dimensions of Personality Munrin Zuckrmuin, James C . Ballenger, and Robert ,% Post I. INDEX

Eye Movement Dysfunctions and Psychosis Philip S . Holzman Peptidergic Regulation of Feeding J . E . Morlgr 7:J . Bartness, B . A. Gosnell, and A. S. Leviiu Calcium and Transmitter Release Ira Cohen and William Van der Kloot

Volume 26

The Endocrinology of the Opioids Mark J. Mzllnn and Albert Hrrz Multiple Synaptic Receptors for Neuroacrive Amino Acid Transmitters-New Vistas Najani A. Shunf

Excitatory Transmitters Related Brain Damage John W. Olney

and

Epilepsy-

Potassium Current in the Squid Giant Axon John R . Clay XHUEX

CONTENTS OF RECENT VOLUMES Volume 28

Biology and Structure of Scrapie Prions Michuel P. McKinley and Stanley B. Prusiner Different Kinds of Acetylcholine Release from the Motor Nerve S. Thesleff Neuroendocrine-Ontogenetic Mechanism of Aging: Toward an Integrated Theory of Aging V. M . Dilman, S. Y. Revsklq, and A. G. Golubev The Interpeduncular Nucleus Barbara J. Morley Biological Aspects of Depression: A Review of the Etiology and Mechanisms of Action and Clinical Assessment of Antidepressants S. I . Ankier and B. E. Leonard Does Receptor-Linked Phosphoinositide Metabollism Provide Messengers Mobilizing Calcium in Nervous Tissue? John N . Hawthorne Short-Term and Long-Term Plasticity and Physiological Differentiation of Crustacean Motor Synapses H . L. Atwood and J. M. Wojtowicz Immunology and Molecular Biology of the Cholinesterases: Current Results and Prospects Stephen Brimijoin and Zoltan Rakoncmy

367

Neurotoxin-Binding Site on the Acetylcholine Receptor Thomas L. Lentz and Paul T. Wilson Calcium and Sedative-Hypnotic Drug Actions Peter L. Carlen and Peter H. W u Pathobiology of Neuronal Storage Disease Steven V. Walkley Thalamic Amnesia: Clinical and Experimental Aspects Stephen G. Waxman Critical Notes on the Specificity of Drugs in the Study of Metabolism and Functions of Brain Monoamines S. Garattini and T. Mennini Retinal Transplants and Optic Nerve Bridges: Possible Strategies for Visual Recovery as a Result of Trauma or Disease James E. Turner, Jeny R. Blair, Magdalene Seiler, Robert Aramant, Thomas W. Laedtke, E. Thomas Chappell, and Lauren Clarkton Schizophrenia: Instability in Norepinephrine, Serotonin, and y-Aminobutyric Acid Systems Joel Gelernter and Daniel P. van Kammen INDEX

Volume 30

Biochemistry of Nicotinic Acetylcholine Receptors in the Vertebrate Brain Jakob Schmidt

INDEX

Volume 29

Molecular Genetics of Duchenne and Becker Muscular Dystrophy Ronald G. Worton and Arthur H . M. Burghes Batrachotoxin: A Window on the Allosteric Nature of the Voltage-Sensitive Sodium Channel George B. Brown

The Neurobiology of N-Acetylaspartylglutamate Randy D. Blakel~and Joseph T. Coyle

Neuropeptide-Processing, -Converting, and -Inactivating Enzymes in Human Cerebrospinal Fluid Lars Terenius and Fred Nyberg Targeting Drugs and Toxins to the Brain: Magic Bullets Lance L. Simpson Neuron-Glia Interrelations Antoniu Vernadakis

368

CONTENTS OF RECENT VOLUMES

Cerebral Activity and Behavior: Control by Central Cholinergic and Serotonergic Systems C. H . Vandenuolf INDEX

Volume 31

Animal Models of Parkinsonism Using Selective Neurotoxins: Clinical and Basic Implications MECIUIPI.J. Zigmond and Edward M.Strickr Regulation of Choline Acetyltransferase Paul M , Salvaterru and James E. Vaughn Neurobiology of Zinc and Zinc-Containing Neurons Chktoplwr J . Frederickson Dopamine Receptor Subtypes and Arousal Ennio Ongtnz and Vincenzo G . Long0 Regulation of Brain Atrial Natriuretic Peptide and Angiotensin Receptors: Quantitative Autoradiographic Studies Juan M . Saauedra, Eero Cmtrhn, Jorge S. Gutkind, and Adil J . Naurali Schizophrenia, Affective Psychoses, and Other Disorders Treated with Neuroleptic Drugs: The Enigma of Tardive Dyskinesia, Its Neurobiological Determinants, and the Conflict of Paradigms John L. Waddington

Volume 32

On the Contribution of Mathematical Models to the Understanding of Neurotransmitter Release H . P a m , I. Parna~,and L. A. &gel Single-Channel Studies of Glutamate Receptors M . S. P. Sawom and P. N . X. Usherwood Coinjection of Xenopus Oocytes with cDNAProduced and Native mRNAs: A Molecular Biological Approach to the Tissue-Specific Processing of Human Cholinesterases Shlonw Sedman and H e n o n u Soreq Potential Neurotrophic Factors in the Marnmalian Central Nervous System: Functional Significance in the Developing and Aging Brain Dalia M . Araujo, Jean-Guy Chabot, and Rhmi Quirion Myasthenia Gravis: Prototype of the Antireceptor Autoimmune Diseases Simone Schonbeck, Susanne Chrestel, and Reinhard Hohlfeld Presynaptic Effects of Toxins AIan L. Haruty Mechanisms o f Chemosensory Transduction in Taste Cells Mylps H. Akabas Quinoxalinediones as Excitatory Amino Acid Antagonists in the Vertebrate Central Nervous System Slpphen N . Devies and Graham L. Collingndge

Nerve Blood Flow and Oxygen Delivery in Normal, Diabetic, and Ischemic Neuropath y Phillip A. Low, Terrence D. Lagerlund, and Philip G . McManis

Acquired Immune Deficiency Syndrome and the Developing Nervous System D o u g h E. Brenneman, Susan K. McCune, and Illanu Gores

IKDEX

INDEX